(2015) Biology of Foodborne Parasites

Food Microbiology Series
Lihua Xiao • Una Ryan
Yaoyu Feng
Series Editor
Dongyou Liu
Biology of Foodborne Parasites, edited by Lihua Xiao, Una Ryan, and Yaoyu Feng (2015)
Lihua Xiao • Una Ryan
Yaoyu Feng
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Preface...................................................................................................................................................... vii
Editors........................................................................................................................................................ ix
Contributors............................................................................................................................................... xi
Section I
1. Introduction and Public Health Importance of Foodborne Parasites......................................... 3
Ronald Fayer
2. Molecular Biological Techniques in Studies of Foodborne Parasites........................................ 21
Una Ryan, Yaoyu Feng, and Lihua Xiao
3. Detection of Parasites in Foods.......................................................................................................41
Ynes R. Ortega and Joan M. Shields
Section II
Important Foodborne Protists
4. Blastocystis....................................................................................................................................... 53
Christen Rune Stensvold
5. Cryptosporidium.............................................................................................................................. 77
Lihua Xiao and Una Ryan
6. Cyclospora cayetanensis.................................................................................................................. 97
Ynes R. Ortega and Jeevan B. Sherchand
7. Cystoisospora...................................................................................................................................111
Jorge Néstor Velásquez and Silvana Carnevale
8. Entamoeba.......................................................................................................................................131
Sandipan Ganguly
9. Enterocytozoon bieneusi.................................................................................................................149
Mónica Santín-Durán
10. Giardia............................................................................................................................................175
Simone M. Cacciò and Marco Lalle
11. Sarcocystis...................................................................................................................................... 195
Benjamin Rosenthal
12. Toxoplasma gondii......................................................................................................................... 209
Dolores E. Hill and Jitender P. Dubey
13. Trypanosoma cruzi......................................................................................................................... 223
Karen Signori Pereira, Flávio Luís Schmidt, Rodrigo Labello Barbosa,
and Luiz Augusto Corrêa Passos
Section III
Important Foodborne Helminths
14. Angiostrongylus............................................................................................................................. 235
Santhosh Puthiyakunnon and Xiaoguang Chen
15. Anisakis.......................................................................................................................................... 255
Simonetta Mattiucci, Michela Paoletti, Paolo Cipriani, Stephen C. Webb,
and Giuseppe Nascetti
16. Clonorchis, Opisthorchis, and Metorchis.................................................................................... 275
Paiboon Sithithaworn, Weerachai Saijuntha, Ross H. Andrews, and Trevor N. Petney
17. Diphyllobothrium, Diplogonoporus, and Spirometra................................................................. 299
Roman Kuchta, Tomáš Scholz, Jan Brabec, and Barbara Narduzzi-Wicht
18. Echinococcus................................................................................................................................. 327
Donald P. McManus
19. Echinostomes..................................................................................................................................351
Jong-Yil Chai
20. Fasciola and Fasciolopsis..............................................................................................................371
Santiago Mas-Coma, M. Adela Valero, and M. Dolores Bargues
21. Gnathostoma.................................................................................................................................. 405
Yukifumi Nawa, Wanchai Maleewong, Pewpan M. Intapan, and Sylvia Páz Díaz-Camacho
22. Metagonimus.................................................................................................................................. 427
Jong-Yil Chai
23. Paragonimus.................................................................................................................................. 445
Yoon Kong, Pham Ngoc Doanh, and Yukifumi Nawa
24. Taenia............................................................................................................................................. 463
Marcello Otake Sato, Caris Maroni Nunes, Megumi Sato, and Jitra Waikagul
25. Trichinella...................................................................................................................................... 481
Xue Bai, Xiaolei Liu, Xiuping Wu, and Mingyuan Liu
Parasites such as helminths and protists are the main pathogens responsible for foodborne illnesses in
both industrialized nations and developing countries. Although funding for research on these pathogens is shadowed by foodborne bacteria and viruses, they have gained increasing attention in both the
research community and public health agencies in recent years. This is largely due to the realization of
the importance of global trade in foodborne disease occurrence, more wide use of culture-independent
diagnostic assays in pathogen detection, and increased use of next-generation sequencing and omics
tools in pathogen characterizations. As a result, significant progresses have been made on biology of
foodborne parasites. This book attempts to showcase some of the most recent developments in research
on foodborne parasites.
The chapters in this book are written by scientists who are actively engaged in research on biology
of major foodborne parasites. Efforts are made to include frontline experts from countries where foodborne parasites exert their heaviest disease burdens. Section I discusses the importance of foodborne
parasites and some key techniques that have been applied to foodborne parasitic pathogen research,
whereas Sections II and III cover individual foodborne protists and helminths, with each chapter providing a state-of-the-art review of biology and pathogenesis of major foodborne parasitic pathogens and
key techniques applicable to pathogen detection and characterization. We hope this book will serve as a
useful guide to medical, veterinary, and food laboratory scientists in need of advanced knowledge and
techniques about foodborne parasites, and a handy textbook for undergraduate and graduate students in
medical, veterinary, and food microbiology.
We thank the authors for their excellent contributions to this book. We also thank the series editor
Dongyou Liu for guidance and the CRC Press senior editor Stephen M. Zollo and his staff members
for the assistance in this project. We appreciate the support and understanding from our colleagues and
families, without whom the work would have been impossible to complete.
Lihua Xiao received his veterinary education in China and his PhD
and postdoctoral training in parasitology in the United States. He joined
the Centers for Disease Control and Prevention (CDC) in 1993, first as
a guest researcher and then as a senior staff fellow. He is currently a
senior scientist in the Division of Foodborne, Waterborne, and
Environmental Diseases, National Center for Emerging and Zoonotic
Infectious Diseases, CDC, Atlanta, Georgia. His earlier research interests were mostly in the immunopathogenesis of malaria and HIV. For
the last 20 years, he has focused mostly on the diagnosis, molecular
epidemiology, comparative genomics, and ecology of foodborne,
waterborne, and zoonotic parasites and is mostly noted for his work on
the genus Cryptosporidium. He has published over 300 original papers
and invited reviews and book chapters, has edited several books and special publications, and is an editorial
board member of several prominent journals in microbiology and parasitology. He holds adjunct faculty
positions at Cornell University, Murdoch University, and East China University of Science and Technology.
He was the 2012 recipient of the Henry Baldwin Ward Medal from the American Society of Parasitologists.
Una Ryan received her undergraduate degree in zoology from
University College Dublin in Ireland in 1988 and her PhD in parasitology from Murdoch University in Western Australia in 1996.
She worked at Murdoch University as a research fellow and became
a tenured member of staff in 2001. She is currently professor in
biochemistry in the School of Veterinary and Life Sciences at
Murdoch University. Her research has focused on the molecular
detection and characterization of waterborne and bloodborne parasites, particularly Cryptosporidium. She has published over 200
original papers and invited reviews and book chapters and is a
specialist editor for Experimental Parasitology. In 2000, she was
awarded the Australian Prime Minister’s Prize for Achievement in
Life Sciences, and in 2014, she received the Bancroft–Mackerras Medal from the Australian Society
for Parasitology for excellence in research.
Yaoyu Feng received her BSc, MSc, and PhD in microbiology in
China and postdoctoral training in water and food safety in the United
States and Singapore. She worked at Tianjin University, Tongji
University, and National Institute of Parasitic Diseases, Chinese
Centers for Disease Control and Prevention. She is currently a professor in microbiology, School of Resources and Environmental
Engineering, East China University of Science and Technology. Since
2000, she has been working on water and food microbial safety,
focusing on molecular biology, epidemiology, diagnosis, taxonomy,
environmental biology, and pathogenesis of Cryptosporidium,
Giardia, microsporidia, and other protozoan parasites of humans and
animals. She has published nearly a hundred papers in international
journals and is an editorial member of three international journals. She received the Distinguished
Young Scholars award from the National Natural Science Foundation of China in 2014.
Ross H. Andrews
Faculty of Medicine
Department of Parasitology
Liver Fluke and Cholangiocarcinoma Research
Cholangiocarcinoma Screening & Care Program
Khon Kaen University
Khon Kaen, Thailand
Faculty of Medicine
Imperial College London
London, United Kingdom
Xue Bai
Key Laboratory for Zoonoses Research
Institute of Zoonosis
Jilin University
Changchun, Jilin, People’s Republic of China
Rodrigo Labello Barbosa
Departamento de Parasitologia
Instituto de Biologia
Universidade Estadual de Campinas
Campinas, Brazil
M. Dolores Bargues
Facultad de Farmacia
Departamento de Parasitologia
Universidad de Valencia
Valencia, Spain
Jan Brabec
Institute of Parasitology
Biology Centre of the Czech Academy of
České Budějovice, Czech Republic
Simone M. Cacciò
Department of Infectious Parasitic and
Immunomediated Diseases
Istituto Superiore di Sanità
Rome, Italy
Silvana Carnevale
Department of Parasitology
National Institute of Infectious Diseases
National Administration of Laboratories and Health
Institutes (ANLIS) “Dr. Carlos G. Malbrán”
Buenos Aires, Argentina
Jong-Yil Chai
Department of Parasitology and Tropical
College of Medicine
Seoul National University
Seoul, South Korea
Xiaoguang Chen
Key Laboratory of Prevention and Control for
Emerging Infectious Diseases
Department of Pathogen Biology
School of Public Health and Tropical Medicine
Southern Medical University
Guangzhou, Guangdong, People’s Republic of
Paolo Cipriani
Department of Public Health and Infectious
Section of Parasitology
Sapienza–University of Rome
Rome, Italy
Sylvia Páz Díaz-Camacho
Facultado de Ciencias y Chimicobiologicas
Universidad Autonoma de Sinaloa
Sinaloa, Mexico
Pham Ngoc Doanh
Institute of Ecology and Biological Resources
Vietnam Academy of Science and Technology
Cau Giay, Vietnam
Jitender P. Dubey
Animal Parasitic Diseases Laboratory
Beltsville Agricultural Research Center
Beltsville, Maryland
Ronald Fayer
Environmental Microbial and Food Safety
Beltsville Agricultural Research Center
Beltsville, Maryland
Xiaolei Liu
Key Laboratory for Zoonoses Research
Institute of Zoonosis
Jilin University
Changchun, Jilin, People’s Republic of China
Yaoyu Feng
School of Resource and Environmental
East China University of Science and Technology
Xuhui, Shanghai, People’s Republic of China
Wanchai Maleewong
Faculty of Medicine
Department of Parasitology
Khon Kaen University
Khon Kaen, Thailand
Sandipan Ganguly
Division of Parasitology
National Institute of Cholera and Enteric Diseases
Indian Council of Medical Research
Kolkata, West Bengal, India
Dolores E. Hill
Animal Parasitic Diseases Laboratory
Beltsville Agricultural Research Center
Beltsville, Maryland
Pewpan M. Intapan
Faculty of Medicine
Department of Parasitology
Khon Kaen University
Khon Kaen, Thailand
Yoon Kong
Department of Molecular Parasitology
Samsung Biomedical Research Institute
Center for Molecular Medicine
Sungkyunkwan University School of Medicine
Suwon, South Korea
Roman Kuchta
Institute of Parasitology
Biology Centre of the Czech Academy of
České Budějovice, Czech Republic
Marco Lalle
Department of Infectious
Parasitic and Immunomediated Diseases
Istituto Superiore di Sanità
Rome, Italy
Mingyuan Liu
Key Laboratory for Zoonoses Research
Institute of Zoonosis
Jilin University
Changchun, Jilin, People’s Republic of China
Santiago Mas-Coma
Facultad de Farmacia
Departamento de Parasitologia
Universidad de Valencia
Valencia, Spain
Simonetta Mattiucci
Section of Parasitology
Department of Public Health and Infectious
Sapienza–University of Rome
Rome, Italy
Donald P. McManus
Infectious Diseases Division
Molecular Parasitology Laboratory
QIMR Berghofer Medical Research Institute
Brisbane, Queensland, Australia
Barbara Narduzzi-Wicht
Istituto Cantonale di Microbiologia
Bellinzona, Switzerland
Giuseppe Nascetti
Department of Ecological and Biological
Tuscia University
Viterbo, Italy
Yukifumi Nawa
Faculty of Medicine
Khon Kaen University
Khon Kaen, Thailand
Caris Maroni Nunes
Faculty of Veterinary Medicine
Universidade Estadual Paulista
Araçatuba, Brazil
Ynes R. Ortega
Center for Food Safety
University of Georgia
Griffin, Georgia
Michela Paoletti
Department of Ecological and Biological
Tuscia University
Viterbo, Italy
Luiz Augusto Corrêa Passos
Centro Multidisciplinar para Investigação
Biológica na Área da Ciência em Animais de
Universidade Estadual de Campinas
Campinas, Brazil
Karen Signori Pereira
Departamento de Engenharia Bioquímica Escola
de Química
Centro de Tecnologia
Universidade Federal do Rio de Janeiro
Rio de Janeiro, Brazil
Trevor N. Petney
Department of Ecology and Parasitology
Institute of Zoology
Karlsruhe Institute of Technology
Karlsruhe, Germany
Santhosh Puthiyakunnon
Key Laboratory of Prevention and Control for
Emerging Infectious Diseases
Department of Pathogen Biology
School of Public Health and Tropical Medicine
Southern Medical University
Guangzhou, Guangdong, People’s Republic of China
Benjamin Rosenthal
Animal Parasitic Diseases Laboratories
Beltsville Agricultural Research Center
Beltsville, Maryland
Mónica Santín-Durán
Environmental Microbial and Food Safety
Beltsville Agricultural Research Center
Beltsville, Maryland
Marcello Otake Sato
Bioterium and Animal Research Laboratory
School of Medicine
Federal University of Tocantins
Palmas, Brazil
Megumi Sato
Faculty of Medicine
School of Health Sciences
Niigata University
Niigata, Japan
Flávio Luís Schmidt
Faculdade de Engenharia de Alimentos
Departamento de Tecnologia de Alimentos
Universidade Estadual de Campinas
Campinas, Brazil
Tomáš Scholz
Institute of Parasitology
Biology Centre of the Czech Academy of Sciences
České Budějovice, Czech Republic
Jeevan B. Sherchand
Public Health Research Laboratory
Department of Medical Microbiology
Tribhuvan University Teaching Hospital
Institute of Medicine
Kathmandu, Nepal
Joan M. Shields
Center for Food Safety and Applied Nutrition
Food and Drug Administration
Laurel, Maryland
Una Ryan
School of Veterinary and Life Sciences
Murdoch University
Murdoch, Western Australia, Australia
Paiboon Sithithaworn
Faculty of Medicine
Department of Parasitology
Liver Fluke and Cholangiocarcinoma Research
Khon Kaen University
Khon Kaen, Thailand
Weerachai Saijuntha
Walai Rukhavej Botanical Research Institute
Mahasarakham University
Maha Sarakham, Thailand
Christen Rune Stensvold
Department of Microbiology and Infection Control
Statens Serum Institute
Copenhagen S, Denmark
M. Adela Valero
Facultad de Farmacia
Departamento de Parasitologia
Universidad de Valencia
Valencia, Spain
Jorge Néstor Velásquez
Department of HIV/AIDS
Francisco Javier Muñiz Infectious Diseases
Buenos Aires, Argentina
Jitra Waikagul
Faculty of Tropical Medicine
Department of Helminthology
Mahidol University
Bangkok, Thailand
Stephen C. Webb
Cawthron Institute
Nelson, New Zealand
Xiuping Wu
National Institute of Parasitic Diseases
Chinese Center for Disease Control and
Shanghai, People’s Republic of China
Lihua Xiao
Division of Foodborne, Waterborne and
Environmental Diseases
National Center for Emerging and Zoonotic
Infectious Diseases
Centers for Disease Control and Prevention
Atlanta, Georgia
Section I
Introduction and Public Health
Importance of Foodborne Parasites
Ronald Fayer
1.1 Disease Burdens of Foodborne Parasites.......................................................................................... 3
1.2 Global Ranking of Foodborne Parasites........................................................................................... 4
1.3 Identifying Parasites and Parasite Burden in Foods......................................................................... 5
1.4 Waterborne Parasites......................................................................................................................... 8
1.5 Fresh Produce–Associated Parasite.................................................................................................. 9
1.6 Parasites in Molluskan Shellfish, Crustaceans, Fish, Frog Legs, and Reptiles.............................. 10
1.7 Meatborne Parasites........................................................................................................................ 13
1.8 Control and Prevention of Foodborne Illness from Parasites..........................................................14
1.9 Future Directions and Trends..........................................................................................................16
1.1 Disease Burdens of Foodborne Parasites
Foodborne illnesses are usually infectious or toxic in nature, and everyone, whether in industrialized or
developing countries, is at risk of acquiring a foodborne illness. The global incidence of foodborne illness
is difficult to estimate, but in 2005, 1.8 million people died from diarrheal diseases, a large percentage of
which can be attributed to contaminated food and drinking water.1 In developed countries, up to 30% of the
population suffers from a foodborne illness each year.1 Less well or poorly documented are the populations
in developing countries that suffer from a wide range of foodborne diseases, including those caused by parasites, but the high prevalence of diarrheal illness in these countries suggests major food safety problems.
In the United States, there were an estimated 48 million annual cases of domestically acquired foodborne illnesses resulting in gastroenteritis, based mostly on data from 2000 to 2008 and the population in
2006 (299 million persons).2 Of these, 9.4 million illnesses were caused by 31 known pathogens including 57,616, 11,407, 76,840, 86,686, and 156 persons for Cryptosporidium spp., Cyclospora cayetanensis,
Giardia duodenalis, Toxoplasma gondii, and Trichinella spp., respectively.3 Another study estimated
the health burden of 14 major pathogens in association with 12 broad categories of food.4 Based on data
from the aforementioned study, within those top 14 pathogens, the annual cost for foodborne illnesses
was $47.2 million for Cryptosporidium, $2 million for Cyclospora, and $2973 million for Toxoplasma.4
Across all 14 pathogens, poultry, pork, produce, and complex foods were responsible for nearly 60% of
the total cost of illness and loss of quality-adjusted life years (QALYs).
An estimated 750 million to a billion people are at risk of infections with foodborne trematodes,
­particularly in Southeast Asia and the Western Pacific region.5,6 Those parasites are the liver flukes such
as Clonorchis sinensis, Fasciola gigantica, F. hepatica, Opisthorchis felineus, and O. ­viverrini, the lung
flukes such as Paragonimus spp., and the intestinal flukes such as Echinostoma spp., Fasciolopsis buski, and
the heterophyids.5 The Food and Agriculture Organization and the World Health Organization estimated
that over 18 million people were actually infected with these trematodes in 2002.7 In 2005, >56 million
Biology of Foodborne Parasites
people were infected with foodborne trematodes, with nearly 8 million suffering severe sequelae and 7158
who died.8 Another study in 2005 estimated that 601.0, 293.8, 91.1, and 79.8 million people were at risk for
infection with only four of these parasites: C. sinensis, Paragonimus spp., Fasciola spp., and Opisthorchis
spp., respectively, and that persons living near freshwater bodies had a 2.15-fold higher risk for infections
than those living farther from the water.6
1.2 Global Ranking of Foodborne Parasites
A global ranking of foodborne parasites with public health issues constituting 85% of the weighted ranking was recently developed.9 Those foodborne parasites considered to be globally important and their
primary food source(s) follow in descending order: Taenia solium (pork), Echinococcus granulosus
(fresh produce), E. multilocularis (fresh produce), T. gondii (meat from small ruminants, pork, beef,
and game meat [red meat and organs]), Cryptosporidium spp. (fresh produce, fruit juice, and milk),
Entamoeba histolytica (fresh produce), Trichinella spiralis (pork), Opisthorchiidae (freshwater fish),
Ascaris spp. (fresh produce), Trypanosoma cruzi (fruit juices), G. duodenalis (fresh produce), Fasciola
spp. (fresh produce aquatic plants), C. cayetanensis (berries and fresh produce), Paragonimus spp.
(freshwater crustaceans), Trichuris trichiura (fresh produce), Trichinella spp. (game meat—wild boar,
crocodile, bear, w
­ alrus, etc.), Anisakidae (saltwater fish, crustaceans, and cephalopods), Balantidium
coli (fresh produce), Taenia saginata (beef), Toxocara spp. (fresh produce), Sarcocystis spp. (beef and
pork), Heterophyidae (fresh and brackish water fish), Diphyllobothriidae (freshwater/saltwater fish), and
Spirometra spp. (fish/reptiles/amphibians) (Figure 1.1). When the parasites were ranked on different
Taenia solium
Echinococcus granulosus
Echinococcus multilocularis
Toxoplasma gondii
Cryptosporidium spp.
Entamoeba histolytica
Trichinella spiralis
Ascaris spp.
Trypanosoma cruzi
Giardia duodenalis
Fasciola spp.
Cyclospora cayetanensis
Paragonimus spp.
Trichuris trichiura
Trichinella spp.*
Balantidium coli
Taenia saginata
Toxocara spp.
Sarcocystis spp.
Spirometra spp.
Normalized overall score
FIGURE 1.1 Global ranking of foodborne parasites (Note: Trichinella spp.* includes Trichinella species except
T. ­spiralis). (From FAO and WHO, Multicriteria-based ranking for risk management of foodborne parasites, Report of
a Joint FAO/WHO Expert Meeting, September 3–7, 2012, FAO Headquarters, Rome, Italy, 2014.
Introduction and Public Health Importance of Foodborne Parasites
weighted criteria, the ranking changed. For example, when ranked on trade criterion scores, the order of
importance changed with T. spiralis, T. solium, T. saginata, Anisakidae, and C. cayetanensis as the top
five. The design of this global ranking process is such that it can be adapted and run by others at national
or regional levels using data more specific to a country or region.
1.3 Identifying Parasites and Parasite Burden in Foods
Identifying parasites in foods or elsewhere in the environment is difficult, requiring different methods than those used for bacterial or viral pathogens. Unlike most other pathogenic microorganisms, most parasitic protozoa and helminths have been difficult or impossible to grow in vitro, and
consequently, their numbers cannot be expanded to facilitate detection and identification, limiting
all identification and biological studies to the initial number of organisms acquired from samples.
Microscopy, the classical tool for identification, has often led to limited sensitivity, high labor input,
and the need for highly specialized technical personnel familiar with these organisms. The way
modern laboratories identify parasites in human, animal, and environmental samples is being revolutionized by molecular methods. The industry associated with methods development has grown
quickly and is now providing virtually all items needed to conduct the polymerase chain reaction
(PCR) technique, the highly accurate, sensitive, and rapid standard for many diagnostic tests. PCR
targets can now be compared to databases of gene sequences determined from clinical and basic
research studies of parasites worldwide. The present methodology in which one PCR assay identifies
one pathogen is evolving to methods like multiplexing that target multiple potential target organisms,
thereby reducing the diagnostic time. Although gene sequencing can provide specific detail regarding foodborne contaminants, interpretation of data requires time and highly skilled personnel, which
are not readily available in many locations. To overcome these obstacles, there is a need to develop
more user-friendly software to facilitate this part of the detection process.
Despite these technical advances, for several reasons, it is difficult to determine the burden of
foodborne parasitic diseases in humans, even within those countries that routinely record human illnesses. First of all, there are little or no requisite reporting systems for the majority of parasitic infections and many differences are found among countries. In the United States in 2011, the Centers for
Disease Control’s list of nationally notifiable infectious conditions contained only four waterborne and
foodborne parasites: Cryptosporidium, Cyclospora, Giardia, and Trichinella.10 In the Netherlands,
Cryptosporidium infection is not a notifiable disease. In the United Kingdom, Cryptosporidium surveillance began in the 1990s, and results from positive laboratory tests are notifiable. In Germany,
Cryptosporidium parvum infection was first notifiable in 2001, but most reports simply indicate the
genus name. Secondly, there are no uniform laboratory standards practices worldwide for the detection
and identification of foodborne parasites, although progress is being made for developing standards for
two parasites—Cryptosporidium and Giardia—but only on fresh leafy greens and berries. Some countries lack both sufficient diagnostic capabilities and reporting capabilities. Furthermore, there is the
impression in many industrialized countries that the prevalence of parasitic infections is low because
of high hygienic standards, such as clean water, sanitary garbage disposal, and sewage processing, and
that contamination of food and drink has been further minimized by quality control standards adopted
by private industry for the production and inspection of meat, fish, some fresh produce, and bottled
drinks. Nevertheless, millions of humans all over the world acquire, through ingestion of contaminated
food and water, a wide variety of protist and helminth infections. As long as population growth continues to increase, clean water resources become more limited, and the volume of international trade in
food continues to expand the prevalence of foodborne and waterborne parasitic infections and can be
expected to remain high and likely increase in some areas.
The most commonly found of these foodborne and waterborne parasites of humans are discussed in
the following chapters of this book and summarized in Table 1.1. The name of the parasite, sources of
infection, the infective stage, clinical signs of infection, methods of diagnosis and detection, and treatments are emphasized. The bases for selecting these parasites include their presence in food and water,
their high prevalence of human infection, the severity of illness they cause, and the recognition that
Biology of Foodborne Parasites
Parasites and Food Associations
Primary Food
Primary Food
Secondary Food
Blastocystis sp.
Cryptosporidium spp.
Water and plants Fresh produce, apple
juice, milk
Possibly molluskan
C. cayetanensis
C. belli
E. histolytica
Water and plants Unknown
Fresh produce
Enterocytozoon bieneusi
G. duodenalis
Water and plants Fresh produce
Sarcocystis spp.
Land animals
T. gondii
Land animals
Meat from cattle,
Fresh produce,
sheep, pigs, chickens, seafood, dairy
goats, game animals
products, water
T. cruzi
Fruit juice
Fresh produce, berries
Angiostrongylus spp.
Aquatic animals Snails, slugs,
and plants
freshwater shrimp,
crabs, frogs, planaria
Aquatic animals Marine fish,
Clonorchis, Opisthorchis, Aquatic animals Freshwater fish
Diphyllobothrium latum Aquatic animals Fish: marine and
Animals and
Amphibians, reptiles,
fish, water
Echinostoma spp.
F. hepatica
F. buski
Aquatic animals Freshwater fish, frogs,
mussels, snails,
snakes, tadpoles
Plants, water,
Watercress or other
land animals
contaminated water
Water plants
Possibly molluskan
Fresh produce
Global Food Attribution
Food associated, proportion
unknown; waterborne and
foodborne outbreaks
documented; water may
be the most important.
Mostly foodborne; basil,
lettuce, berries.
Food associated, proportion
unknown; waterborne
route and food handlers
Food associated, proportion
unknown; waterborne and
foodborne outbreaks
documented; food
handlers implicated.
S. hominis and
S. suihominis meatborne.
Multiple routes of
infection; transmission via
meat >20%; waterborne
outbreaks documented.
Foodborne outbreaks
documented in limited
geographic area from
insect contamination.
All foodborne.
All foodborne.
All foodborne.
All foodborne.
Possibly pork or meat
from other infected
land animals
All food- and waterborne.
All foodborne.
Raw sheep and goat
All food- and waterborne.
All foodborne.
(Continued )
Introduction and Public Health Importance of Foodborne Parasites
TABLE 1.1 (Continued )
Parasites and Food Associations
Primary Food
Gnathostoma spp.
Haplorchis and
Metagonimus yokogawai
P. westermani
T. solium
T. saginata
Trichinella spp.
Primary Food
Secondary Food
Freshwater fish,
Possibly water
freshwater eels, frogs, containing infected
birds, and reptiles
water fleas
Aquatic animals Freshwater and
brackish water fish
Aquatic animals Freshwater and
brackish water fish
Aquatic animals Freshwater crustaceans Boar meat
Land animals
Land animals
Land animals
Horsemeat, game meat
Global Food Attribution
All foodborne.
All foodborne.
All foodborne.
All foodborne.
All meatborne.
All meatborne.
All meatborne.
Source: Adapted and modified in parts from FAO and WHO, Multicriteria-based ranking for risk management of foodborne
parasites, Report of a Joint FAO/WHO Expert Meeting, September 3–7, 2012, FAO Headquarters, Rome, Italy,
international transport of fresh food supplies can bring these parasites to places where they were once
rare or unknown. As indicated in Table 1.2, the volume of food products shipped internationally is enormous. The ability to inspect such great quantity and such diversity of food products for the wide range of
potential pathogens is a daunting task, often with insufficient resources.
Some very important parasites of public health concern such as the malaria parasites and those that
cause onchocerciasis and schistosomiasis have not been included because infection via ingestion of
food or water is not considered their normal route of transmission. Still others such as Balantidium,
Ascaris, Trichuris, and Toxocara were not included because the proportion of foodborne transmission is
unknown and thought to be relatively low. Blastocystis sp. and C. belli were included because they are
found worldwide and are emerging in recognition.
International Trade Volumes
Food Source (Edible Products)
Game/wild animals
Shellfish (oysters, clams, mussels)
Crustaceans (freshwater)
Crustaceans (marine)
Fish (freshwater)
Fish (marine)
Vegetables (fresh)
Fruits (fresh, nonberries)
Fruit juice
Total tonnage
2009 or 2010 Trade Volumes (Tons)
Source: Based on FAOSTAT, FAO Fisheries and Aquaculture Statistics Service and on FAO
Statistical databases, 2012,
Biology of Foodborne Parasites
1.4 Waterborne Parasites
Ultimately, humans acquire foodborne and waterborne parasites either from other humans or from animals via fecal contamination of the environment, except for some free-living amoebae and ciliates.11
Waterborne parasites can be ingested directly with drinking water, from accidental ingestion of recreational water, or secondarily from contaminated fresh produce or other foods exposed to contaminated
water during irrigation or washing. Encysted stages of parasites have been identified in surface waters
such as rivers, lakes, ponds, reservoirs, and even tidal and marine waters.
In the United States, the levels of Cryptosporidium, Giardia, and other contaminants in public drinking water are controlled under a series of rules for monitoring them in source water, the most recent
being the Long Term 2 Enhanced Surface Water Treatment Rule (LT2).12 Other countries have similar
rules and organizations responsible for providing safe potable water. However, despite efforts to prevent
transmission of waterborne parasites by such agencies in developed countries, at least 191 outbreaks due
to the waterborne transmission of parasitic protozoa were reported; from 2004 to 2010, 46.7% occurred
on the Australian continent, 30.6% in North America, and 16.5% in Europe.13
Blastocystis sp. (Chapter 4) consists of 14 subtypes, associated with different hosts, and distinguishable
only by molecular methods. This enigmatic parasite belonging to the kingdom Chromista is one of the
most commonly found parasites infecting the intestine of humans and animals worldwide.14 Generally,
developing countries have higher prevalences of Blastocystis infections than developed countries, linked
to consumption of contaminated water or food, poor hygiene, and exposure to animals. Prevalence was
found to be low in developed countries such as Japan (0.5%–1%) and Singapore (3.3%) but high in developing countries such as Argentina (27.2%), Brazil (40.9%), Cuba (38.5%), Egypt (33.3%), and Indonesia
(60%).15 It has been detected by molecular analysis of drinking water sources in Turkey, Thailand,
Malaysia, and Nepal and associated with human infections in close proximity to water sources.15–17
Cyclospora (Chapter 6) is a genus within a large group of protozoa collectively known as coccidia,
most of which possess an environmentally robust oocyst stage capable of remaining infectious in moist
conditions for weeks or months. C. cayetanensis, the only species in the genus that infects humans, is
endemic in countries such as Haiti, Guatemala, Peru, and Nepal and has been detected in water used for
human consumption in the United States, Nepal, and Turkey.18 Three outbreaks thought to be associated
with drinking water have been reported in Lima, Peru and Izmir, Turkey.18 In the United States, epidemiologic studies suggested that tap water in a physicians’ hospital dormitory was the most likely source
of an outbreak of cyclosporiasis and that stagnant water in a storage tank might have contaminated the
tap water after a pump failure.19 However, inability to explain how the storage tank could have become
contaminated renders the waterborne concept questionable.
Cystoisospora (syn. Isospora in mammals) (Chapter 7) is also classified within the large group of protozoa collectively known as coccidia. Cystoisospora belli (syn. Isospora belli) is the only known species
in this genus that infects humans. Found worldwide, poor sanitation and fecal contamination of water
and food are the most likely explanations for the spread of C. belli, although there are no reports that
actually document the presence of this organism in water or food. Outbreaks have been reported from
mental wards, daycare centers, World War II veterans in the Pacific, and especially in AIDS patients in
many parts of the world.20
E. histolytica (Chapter 8) and E. dispar parasitize about 10% of the world population, although 90% of
infections are thought to be asymptomatic; nevertheless, amoebiasis causes up to 110,000 deaths a year.21
It was a nationally notifiable disease in the United States before 1995, and from 1976 to 1985, roughly
3000–7000 cases were reported annually. Transmission is usually by ingestion of cysts in contaminated
water or food contaminated from infected food handlers and from person-to-person contact,22 although
nonhuman primates in close contact with humans can be a source of infection.
G. duodenalis (syn. G. intestinalis; G. lamblia) (Chapter 10) is a flagellated protozoan of the intestine responsible for an estimated 1 billion cases of diarrhea annually. Within G. duodenalis are nine
genotypes, referred to as assemblages A–H and the quenda genotype, which differ from each other
genetically and in host specificity. Only assemblages A and B infect humans. Waterborne infections have
long been recognized among hikers and campers who acquired giardiasis after taking drinking water
Introduction and Public Health Importance of Foodborne Parasites
from streams and ponds. Because beavers were occasionally seen where water was acquired, the name
“beaver fever” was given to the illness. G. duodenalis was the most frequently identified pathogen in
all drinking water outbreaks reported in the United States during 1971–2006, responsible for 121 of 432
outbreaks with an identified etiology.23 In addition, waterborne-associated outbreaks have been reported
in Norway, Finland, Canada, Turkey, Malaysia, New Zealand, and Australia.23 In the United States in
2006–2010, there were over 19,000 cases reported annually from all sources with peak numbers in
children 1–9 years of age and in adults 35–44 years of age during early summer through early fall.24,25
Of the present 25 valid species of Cryptosporidium (Chapter 5), the most prevalent in humans are
C. hominis, C. parvum, and C. ubiquitum. Cryptosporidiosis is estimated to account for 20% of all cases
of childhood diarrhea in developing countries, and globally, the percentage of a population infected at
some time in life has been estimated to range from 20% to 90% depending on location.26 Many outbreaks
of cryptosporidiosis have been related to public drinking water and have affected many thousands of
people in developed countries despite treatment by sedimentation, filtration, and chemical disinfection
that has greatly reduced transmission of viral and bacterial pathogens in drinking water. These outbreaks
characteristically have been associated with flooding or failures at water treatment facilities that have
enabled contaminated water to pass either through the treatment system, as in the massive Milwaukee
cryptosporidiosis outbreak affecting nearly 400,000 people,27 or from persons drinking untreated water.
To compensate for breakdowns at drinking water treatment facilities, “boil water” alerts have been
issued. Irrigation waters have been suggested as a route by which Cryptosporidium contaminates fresh
fruits and vegetables. For example, 36% of the water used to irrigate crops traditionally eaten raw in
the United States and Central America tested positive for C. parvum oocysts, 48% of irrigation waters
examined in Mexico contained Cryptosporidium oocysts, and irrigation waters in Norway were also
found contaminated with C. parvum.26
T. gondii (Chapter 12) is another coccidian parasite found worldwide in humans and known to infect
virtually all vertebrate species. Felids, the only definitive hosts, produce the environmentally robust
oocyst stage. Outbreaks of toxoplasmosis associated with drinking contaminated water have been
reported in Panama, Brazil, French Guiana, and India.13,28,29
Recreational water activities such as swimming and water park activities have been waterborne sources
associated with other outbreaks of cryptosporidiosis, giardiasis, and possibly microsporidiosis.30–33
Of the five species of Echinococcus tapeworms (Chapter 18), two are responsible for most of the morbidity and mortality. E. granulosus causes cystic hydatid disease, affecting an estimated 2–3 ­m illion
people globally. Sheep, cattle, goats, pigs, horse, camels, and other ungulates are typical hosts that
bear the cysts. Dogs usually acquire infection after eating infected liver and lungs from hydatidharboring livestock after owners conduct home or field slaughter. Humans become exposed to the
tapeworm eggs from contact either with an infected dog or from eggs in the environment including
contaminated drinking water. In the northern hemisphere, E. multilocularis causes alveolar hydatid
disease affecting about 0.3–0.5 million people. Foxes and canids, the definitive hosts, become infected
with the adult tapeworm. A wide range of small mammals act as intermediate hosts. Eggs excreted by
definitive hosts are extremely environmentally resistant and can remain viable for months or years.
People become accidental intermediate hosts after ingesting eggs, through either water, contaminated
foods, or contact with infected definitive hosts. In Japan, drinking well water was identified as an
epidemiological risk factor.34
1.5 Fresh Produce–Associated Parasite
Based on data from outbreak-associated illnesses between 1998 and 2008, 46% of 9.6 million estimated
annual foodborne illnesses in the United States were attributed to fresh produce, and more illnesses were
associated with leafy vegetables (23%) than any other commodity.35 Fresh fruits and vegetables worldwide
have been found contaminated with many species of parasites, some of which include Cryptosporidium,
Cyclospora, Giardia, Entamoeba, Fasciola, and Angiostrongylus. Contamination can result from intermediate hosts depositing parasites on the plants; from fertilizing with animal or human feces; from
fecal contamination by wild animals; from irrigation water contaminated with feces; from food handlers
Biology of Foodborne Parasites
during picking, processing, and preparation of foods; or from contaminated rinse water. Even thorough
washing with clean water might not remove all encysted parasites from contaminated fresh produce.35
Many cases of giardiasis have been related to ingestion of drinking and recreational water, but foodborne outbreaks of giardiasis are infrequently reported in the United States. In the decade 2000–2010,
less than 1% of foodborne outbreaks with an identified cause were attributed to Giardia.36 In England,
eating lettuce was associated with increased risk for giardiasis.37 Use of reclaimed wastewater for irrigation is associated with finding Giardia cysts on fresh produce.38 However, foodborne outbreaks of giardiasis have generally been caused by direct contamination by an infected food handler or by animals.39
Oocysts of Cryptosporidium have been detected on many types of fresh produce worldwide. From
studies in North America, Latin America, Europe, and Asia, Cryptosporidium has been detected on
basil, blackberries, cabbage, celery, Chinese cabbage, cilantro, green onions, mung bean sprouts, parsley,
romaine, spinach, water spinach, and in unpasteurized apple cider.25 Some of these products have been
associated with foodborne outbreaks.
Similar products have been contaminated with Cyclospora, and outbreaks of cyclosporiasis in North
America have been associated with ingestion of contaminated raspberries, mesclun lettuce, snow peas,
and basil.40,41 For example, in Florida in 1995, there was an outbreak of cyclosporiasis among 45 residents
who has not recently traveled outside the United States. In 1996, outbreaks were reported in 20 states,
the District of Columbia, and 2 Canadian provinces among 1465 persons who had eaten Guatemalan
raspberries. In 1997, there were 510 cases among persons who had eaten raspberries from Guatemala
or mesclun lettuce. In 1997, there were approximately 185 cases among persons who had eaten foods
containing fresh basil. In 1998, raspberries from Guatemala were banned from import into the United
States and no cases of cyclosporiasis were reported, whereas in Canada, raspberries were imported and
an outbreak occurred.
Recently, the protozoan T. cruzi (Chapter 13) has been found in unpasteurized fruit juices such as
sugarcane juice, guava, and acai berry juice. These juices have been identified as sources of at least 10
outbreaks of Chagas disease.42–44 At night, when acai juice was being made, insects attracted by lights had
fallen into the juice and were ground up in the fruit pulp releasing trypanosome parasites into the mixture.
Clinical signs and serologic data provided strong evidence that an outbreak of eosinophilic meningitis
in Taiwan from Angiostrongylus cantonensis (Chapter 14) was associated with drinking raw vegetable
juice.45 Another outbreak was reported among travelers returning from the Caribbean to the United
States after having eaten lettuce in a Caesar salad.45 Most cases of eosinophilic meningitis in New
Caledonia were thought to result from an influx of infected planarians in local produce gardens and their
subsequent accidental ingestion on lettuce and other raw vegetables.46
F. hepatica (Chapter 20) is a liver fluke infecting perhaps as many as 2.4 million people worldwide
who have ingested contaminated water or leafy green plants such as watercress, lettuce, and spinach.47
Fascioliasis is an endemic problem in parts of South America and is emerging in Southeast Asia. Where
sanitation is poor or nonexistent and in known endemic areas, people are cautioned to eat only thoroughly cooked vegetables.
The role of food in transmission of echinococcosis has not been definitively established. For 40 infected
persons in Germany who had eaten unwashed or uncooked vegetables, salads, herbs, berries, or mushrooms, these products did not appear to be an important risk factor for alveolar hydatid disease.48 Having
eaten unwashed strawberries or having chewed grass was more common among patients than controls, but
calculations suggested that these exposures could at most account for only a quarter of the overall risk for
infection. Drinking water from natural sources had no identifiable association with the disease.
1.6 Parasites in Molluskan Shellfish, Crustaceans, Fish, Frog Legs, and Reptiles
Shellfish and fish are sources of many foodborne parasites worldwide. In 2002, more than 18 million
people were infected with zoonotic trematodes from fish worldwide, although the problem is especially
high in Asia where fish are the main source of protein.49 Two major changes in fish consumption are
taking place. Worldwide consumption of fish per person has nearly doubled in the past four decades;
at the same time, fish capture on a global scale has leveled off because most oceanic food fish have
Introduction and Public Health Importance of Foodborne Parasites
World Fisheries and Aquaculture Production (in Millions of Tons)
Wild caught
World total
Source: Data from FAO World Review of Fisheries and Aquaculture, 2012. (Dec 2, 2014)
been fully captured. In 1970, 5.3% of the fish for human consumption came from aquaculture; in 1980,
it was 9%; in 2000, it was 32.2%; and in 2006, it was 43%, which was 45.5 million tons of farmed fish
versus 60 million tons of freshwater and marine capture fish for human consumption.50 To meet the
market demand, there has been tremendous growth of fish farming (Table 1.3). Of the total global aquaculture production in 2000, 91.3% was farmed in Asia where fish-borne trematodes are endemic.8 For
example, in China, the leading global producer of grass carp, a major intermediate host of foodborne
trematodes, production increased from >10,000 tons in 1950 to >3 million tons in 2002; here, grass carp
is traditionally eaten raw as sushi or yusheng zhou.8 With the expansion of aquaculture, there has been
an increase in intermediate hosts and increased use of wastewater and feces to supplement water availability and provide nutrients. Although this is more common in low-income countries, populations at
risk are increasing because of faster and more numerous transportation systems supporting expanding
international markets. Some other farm-raised fish include Atlantic salmon in the United States, United
Kingdom, Chile, Norway, Canada, Ireland, and Iceland; rainbow trout in the United States and Canada;
tilapia in the United States and China; and catfish in the United States. In North America, catfish and
tilapia are farmed under strict environmental conditions that restrict parasites. However, freshwater
fishes such as walleye and pike and pelagic fish such as herring, mackerel, and salmon are potential
sources of cestode and nematode parasites.
The environmentally rugged oocyst stage of Cryptosporidium and Toxoplasma as well as cysts of
Giardia can survive for months in moist, cool conditions, in ponds, lakes, rivers, and brackish waters
where they can be ingested by and survive within oysters, clams, cockles, and mussels that filter such
small particulates from the water.51–53 However, there are no documented cases of infection with any of
these parasites that have resulted from eating raw shellfish.
Several nematode parasites are transmitted to humans that eat undercooked or raw mollusks, crustaceans, and cold-blooded vertebrate hosts. A. cantonensis (Chapter 14), a rat lungworm endemic in
Southeast Asia, Pacific islands, and Australia, also has been found in rats and humans in Cuba54 and in
rats in Louisiana.55 Larvae excreted in rat feces are eaten by snails or slugs and develop into infective
larvae. When humans eat food contaminated with infective larvae such as slugs or aquatic and terrestrial snails, planarians, freshwater shrimp, land crabs, or frogs, the infective larvae invade intestinal
tissue, pass through the liver and lungs, and reach the eye or possibly the central nervous system where
they can cause death. For example, in Taiwan, the disease is the commonest among children, with the
­highest rates during the summer rainy season, and was found in a child who raised snails as pets; infection is often diagnosed among Thai laborers who ingest raw snails (ProMED-mail Angiostrongylus
­meningitis—Taiwan 20130206.1532259). Another species, A. costaricensis, has been reported in Central
America and can cause a similar disease in humans.56 Infection is difficult to control or prevent in
endemic areas because larvae can survive in drinking water and on vegetables.
Biology of Foodborne Parasites
Lung flukes of the genus Paragonimus (Chapter 23) cause paragonimiasis in humans, cats, rats, mice,
monkeys, dogs, pigs, and other animals after eating freshwater crustaceans such as crabs, prawns, or
crayfish. P. westermani and several other species are found in China, the Philippines, Japan, Vietnam,
South Korea, Taiwan, and Thailand. In Asia, an estimated 80% of freshwater crabs carry P. westermani.1
P. africanus is found in Africa, and P. mexicanus is found in Central and South America. P. westermani, P. kellicotti, and P. mexicanus have been found in the United States in crayfish and crabs.57 Of 34
case reports and 1 survey of infections in the United States, at least 10 were local, more than 20 were
imported, and 9 were campers who had eaten raw crayfish from rivers in Missouri.57,58 P. kellicotti or
other species have caused infection after eating raw freshwater crabs in sushi.
The major genera of concern for anisakid infections in humans are Anisakis and Pseudoterranova
(Chapter 15). Anisakiasis, also known as whale worm, herring worm, sealworm, and codworm disease,
is the result of accidental human infection with the larval stage of several nematodes found in raw or
pickled fish such as herring, mackerel, and salmon whose major definitive hosts are whales and porpoises
and Pseudoterranova larvae in fish such as cod, flounder, and flukes whose major definitive hosts are
seals.59 About 20,000 human cases have been reported, mostly from Japan with the others from the
Netherlands, Germany, France, and Spain.60 Each year, there are about 2000 new cases in Japan, 500 in
Europe, and 50 in the United States.61 Although most anisakid larvae in spring spawning herring, blue
whiting, and mackerel from the Northeast Atlantic were found in the abdominal cavity, the prevalence of
larvae in fillets from these fish was 15%–60%, 32%–77%, and 89%–100%, respectively, raising concern
for food safety.62
Gnathostoma (Chapter 21) are also roundworms especially common in Southeast Asia and Japan but
also in parts of the western hemisphere such as Mexico and in Central and South America.63 Humans
and other mammals become infected after eating raw fish, eels, other cold-blooded vertebrates, and even
birds and pigs with encysted larvae in their muscles.
Intestinal and liver trematodes are major fish-borne zoonoses of humans. In Asian countries where
foodborne trematodes are endemic, the wholesomeness of fish meat constitutes a public health concern
for the safety of local populations and for safe food products exported to other countries.
In China, infections with C. sinensis (the Chinese liver fluke) have more than tripled within a single decade to about 15 million people infected in 2004.3 Most case reports in the United States were
immigrants from China, Japan, Korea, and Southeast Asian countries, although four were native-born
Hawaiians and one was a person who possibly became infected after eating imported fish. Related parasites Opisthorchis and Metorchis (Chapter 16) also encyst under the scales of fish and infect humans
that eat raw or undercooked fish. Clonorchis and Opisthorchis are very similar morphologically and
biochemically. Usually, carp and related fish become infected from eating protobranch snails, not commonly found in the United States.57 Metorchis conjunctus, the Canadian liver fluke, is a parasite of carnivores in North America. In NW Ontario, Canada, 10 persons were infected with Metorchis.64 Nineteen
people who ate raw fish prepared from the white sucker, caught in a river north of Montreal, Canada,
became infected.
Echinostoma (Chapter 19) and Metagonimus (Chapter 22) are genera of very small flukes of only a
few millimeters in length that also infect humans that eat raw fish. Some cestodes (tapeworms) are also
acquired by eating raw or undercooked cold-blooded vertebrates.
The larval stages of Spirometra species (Chapter 17) cause a disease called sparganosis while migrating through the body after persons accidently ingest copepods in water or by eating undercooked
amphibians, reptiles, or fish. Sparganosis has been reported sporadically around the world with a higher
prevalence in Asian countries including South Korea, Japan, Thailand, and China.65 To date, more than
1000 cases of humans have been reported in 25 provinces in mainland China where it is emerging
because of food consumption habits and the practice of treating wounds or other lesions with poultices
of frog or snake flesh.65
Related biologically and morphologically to Spirometra are the many species of Diphyllobothrium
(Chapter 17). D. latum, called by its common names the “broad tapeworm” or “fish tapeworm,” is the
largest tapeworm that infects humans. About 20 million people worldwide are infected, making D. latum
the most important fish-borne zoonoses caused by a cestode.66 Of an estimated 100,000 people infected
with D. latum in North America, most are in the Great Lakes area. The greatest prevalence of infection
Introduction and Public Health Importance of Foodborne Parasites
is in northern Europe, Asia, and North America but also in Uganda and Chile. Fish infected with larva
have been transported to and consumed in countries with no known infected fish such as Brazil. Other
species of Diphyllobothrium infect humans, but less often.
All of the aforementioned parasites are killed by thorough cooking, whereas raw or undercooked
dishes, popular in some venues, or the tasting of fish during preparation of special dishes carries the risk
of infection.
1.7 Meatborne Parasites
Livestock are extremely important worldwide in human nutrition and socioeconomic development,
­contributing around 13% of calories and 28% of protein.67 In developing countries, they are a source
of income and employment, contribute manure and serve as draft animals for crop production, and
represent financial assets. Over the past four decades, growth in population and income have resulted in
greater consumption of animal protein, especially in developing countries, with global meat consumption expected to rise nearly 73% by 2050.67
Based on data from outbreak-associated illnesses between 1998 and 2008, 22% of 9.6 million estimated annual foodborne illnesses in the United States were attributed to meat-poultry commodities
(beef, game, pork, and poultry).36 Meat can be a source of many parasites. Encysted, encapsulated, or
free stages of parasites have been found in the muscles and other tissues. These parasites have specific
life-cycle stages in the animals and can be acquired by humans eating raw or undercooked meat or meat
products. The greatest cause of infection is from ingestion of raw or undercooked meat products attributed to customary social habits and taste preferences. Virtually, all are killed by thorough cooking and
some can also be killed by freezing.
Sarcocystis species (Chapter 11) in the muscles of pigs, cattle, and sheep can be microscopic or macroscopic depending on the species and the definitive host, but infection in human definitive hosts is rarely
reported.68 When S. hominis or S. suihominis from cattle and pigs, respectively, are ingested by humans,
bradyzoites released from the cysts develop into sexual stages in the intestine that fuse and produce
fully developed sporocysts that are excreted, sometimes asymptomatically and at other times, perhaps
depending on the parasite load, with abdominal discomfort.68
Worldwide, it is estimated that over 6 billion people have been infected with T. gondii (Chapter 12).69
Seroprevalence, measured by IgG, has been reported as 6.7% in Korea, 12.3% in China, 23.9% in
Nigeria, 46% in Tanzania, and 47% in rural France with some regions as high as 98%.69 In the United
States, it is estimated that more than a million people become infected with T. gondii annually, including
over 21,000 with ocular lesions (both asymptomatic and symptomatic).70 Furthermore, from about 500
to 5000 newborns acquire congenital toxoplasmosis each year resulting in mental retardation, seizures,
blindness, and death.70
Toxoplasma, capable of infecting virtually all warm-blooded animals, has been identified in muscles
and other organs such as heart, brain, and liver of food animals such as cattle, sheep, pigs, goats, horses,
chickens, and other poultry, as well as many game animals such as bear, deer, elk, moose, walrus,
pheasant, and quail. In a case-control study conducted in the United States, attributable risk factors for
acquiring toxoplasmosis included eating raw ground beef (7%), rare lamb (20%), and locally produced
cured, dried, or smoked meat (22%) and working with meat (5%).71 Nonmeat sources included drinking
unpasteurized goat’s milk (4%), having three or more kittens (10%), and eating raw oysters, clams, or
mussels (16%).71
T. spiralis, the roundworm that causes trichinellosis (Chapter 25), is perhaps the best known of these
meatborne parasites worldwide. Surveys performed by the International Commission on Trichinellosis
identified 3281 cases from 103 outbreaks in 2004 and 2409 cases (including 5 deaths) from 44 outbreaks
in 2005, indicating that trichinellosis was a concern in Romania, Bulgaria, Croatia, Serbia, Poland,
Argentina, China, and Laos where backyard or free-roaming pigs are the sources of human cases.72,73 In
the United States, only 72 cases of trichinellosis were diagnosed between 1997 and 2001 and 66 cases
were diagnosed in 2002–2007.74,75 Other species of Trichinella are found in wildlife and have contributed
to infections after undercooked meat or meat products from bears, boars, crocodiles, walruses, and other
Biology of Foodborne Parasites
species are eaten. Historically, pork has been the major source of infection, but in the United States, T.
spiralis is present in only about 1 hog out of 10,000. This low prevalence is a result of animal husbandry
practices that include the elimination of feeding raw or undercooked garbage containing infected meat,
infected rodents, wildlife, or infected hog carcasses. Most young pigs are fed a combination of soybean
meal and corn until they are sent to slaughter; in some places, hogs are fed mostly potatoes and byproducts, sugar beets, or forage. In countries that conduct meat inspection for trichinae, infection from
pork is rare.
T. solium, “the pork tapeworm,” is distributed worldwide, with higher prevalence rates in less developed countries. Pigs that ingest T. solium eggs from the environment develop the larval cyst stage in
their muscles called a cysticercus. Humans that eat raw or undercooked pork containing cysticerci
develop tapeworms in their intestine. Humans that ingest the eggs of T. solium develop neurocysticercosis when the larval stage locates in the central nervous system. Mexico and Brazil report the highest incidence of neurocysticercosis, but the prevalence in Latin America is poorly documented and
T. saginata, “the beef tapeworm,” is the most prevalent large tapeworm of humans. It can be transmitted by ingestion of the infectious cysticercus stage in measly beef that is partially cooked, smoked, or
pickled, although raw beef is the most common source of infection. This parasite is distributed worldwide with a higher prevalence of human infection in less developed countries and countries where raw
or undercooked beef is eaten often. Its prevalence varies regionally, but it is the most common tapeworm
in humans in central Europe77 and is more common than T. solium in the United States. In the United
States and the European Union countries, the carcasses of animals found to be infected with T. saginata
cysticerci (bladder worms) during meat inspection are downgraded requiring extra handling and freezing or they are condemned if infected.77
Meat inspection for trichinellosis and cysticercosis is required in European Union countries as
described under Regulation 854/2004 for pigs and cattle at slaughter and in six other exporting countries
including Australia, Canada, Japan, New Zealand, Norway, and the United States.
1.8 Control and Prevention of Foodborne Illness from Parasites
Personal preferences and traditions for the preparation and consumption of foods play a role in the
acquisition of parasitic disease. Raw and undercooked foods are sources of several parasitic infections. Whether it is eating raw snails that transmit Angiostrongylus; raw fish that transmit a variety of
trematodes; or raw pork, lamb, beef, horsemeat, or wild game that transmit Toxoplasma, infection with
these and other parasites can be prevented simply by thorough cooking. It may be difficult to overcome
­customary eating traditions, but education of consumers, especially in endemic areas, is a relatively inexpensive and reasonable approach to reduce the spread of foodborne parasitic diseases.
In the United States, agencies responsible for safe drinking water, produce, seafood, and meat include
the United States Environmental Protection Agency (USEPA), the Food and Drug Administration
(FDA), the United States Department of Commerce’s (USDC) National Oceanic and Atmospheric
Administration (NOAA), and the United States Department of Agriculture (USDA). Comparable government agencies exist in many developed countries and in some developing countries.
In the United States, the Office of Groundwater and Drinking Water of the USEPA provides water
treatment plants with regulations on filtration, chemical flocculants and disinfectants, turbidity, and
microbial contaminants. The USEPA also sets rules for water-testing methods and schedules, as well as
techniques for treating contaminated water. The Safe Drinking Water Act established the limits on the
levels of Cryptosporidium, Giardia, and other contaminants in public drinking water. Because contamination with these parasites is a concern, USEPA developed rules for monitoring them in source water.
The most recent is the LT2.12 In England and Wales, the Drinking Water Inspectorate (DWI) assures
that water supplies are safe and drinking water is high quality by scrutinizing water company activities, commissioning research to provide evidence-based data on drinking water, and publishing data
on drinking water quality ( Likewise, in Japan, the Office of Drinking Water
Introduction and Public Health Importance of Foodborne Parasites
Quality Management of the Ministry of Health, Labour, and Welfare has established guidelines for
drinking water safety and quality. Other developed countries have established similar agencies and rules
to provide safe drinking water.
Through the Food Safety Modernization Act, the FDA has authority for enforcement and inspection to
prevent unsafe foods from being distributed, but there is no routine postharvest inspection for microbial
contaminants, or markers for them, by a governmental agency. Industry practices self-compliance by
producers and packers. These operations often take place in field packing sheds, whereas leafy greens
that are put into salad bags are handled by processors who voluntarily have their production and handling
operations such as for leafy greens examined by third-party certification audit programs. Large buyers
such as grocery cooperatives or other national or regional retail outlets require documentation demonstrating compliance with industry-wide food safety production and pre- and postharvest practices before
they purchase from growers and packers/processors to ensure that the produce has been grown and
handled with the best available practices to minimize the risk of contamination and thereby limit their
liability. The FDA produces many guides with recommendations for industry practices, and specialty
crop producer groups have published customized guidance documents. For example, from 1996 to 2008,
82 foodborne illness outbreaks were associated with the consumption of fresh produce of which 34.1%
were linked to the consumption of leafy greens accounting for 949 illnesses and 5 deaths.78 Most of these
outbreaks were caused by Escherichia coli O157:H7, Cyclospora, and Salmonella.78 To improve the
safety of fresh produce, FDA issued their Guide to Minimize Microbial Food Safety Hazards for Fresh
Fruits and Vegetables that provides general food safety guidance where safety might be compromised
during the growing, harvesting, transportation, cooling, packing, and storage of fresh produce.78 It also
alerts fruit and vegetable growers, packers, processors, and shippers to potential microbiological hazards
associated with land history, nearby land use, water quality, worker health and hygiene, equipment cleaning and sanitation, and product transportation.
The USDC Seafood Inspection Program provides lot inspection (audit) and sampling services on
a voluntary, fee-for-service basis. These inspection services can determine adherence to minimum
acceptable quality (MAQ); US Grade A, B, or C attributes; and buyer specifications such as net weight,
size, count, and/or other product attributes as defined by the buyer.79 These services conform to global
activities to harmonize inspection protocols and are designed to enhance the safety, wholesomeness,
economic integrity, and quality of seafood available to consumers.79 If there are no buyer specifications, the minimum inspection effort applied to every lot inspection is adherence to the MAQ standard
of quality and condition, flavor, and odor.79 As aquaculture production increases, products should be
monitored carefully to exclude sources of parasites especially in Asian countries where fish-borne
trematodes are endemic.
All interstate food processors of poultry and red meats are inspected by the USDA’s Food Safety
Inspection Service (FSIS). In 1997, the federal government introduced a food safety regulation for all
federal- and state-inspected meat and poultry slaughter and processing plants as well as foreign plants that
export to the United States called the hazard analysis and critical control point (HACCP) rule. As an example of the scope of inspection responsibilities, in 2012, FSIS inspectors examined 32.4 M cattle, 112.3 M
hogs, 2.0 M sheep and lambs, 8576.2 M chickens, and 250 M turkeys (
However, the Federal Meat Inspection Act (Title 21—Food and Drugs, Chapter 12—Meat Inspection)
contains no specific information on pathogens, only inspection checks for product adulteration, sanitation
practices, and misbranding. There is no mandatory inspection of meat for T. gondii or T. spiralis in the
United States. Voluntary inspection of pork by the pooled sample digestion method for T. spiralis is allowed
under the Code of Federal Regulations Title 9, Chapter 3 (9 CFR § 318.10) and the title “Prescribed treatment of pork and pork products to destroy trichinae.” FSIS Directive 6100.2 lists specific disease conditions
for inspection of carcasses including cysticercosis (9 CFR 325.7) in beef, pork, and lamb and Sarcocystis
in carcasses in general.80 European Union countries and Russia require inspection for Trichinella in pork
exported from the United States. Requirements can be found at
This level of regulation and inspection does not exist in most developing countries. Local and homeslaughtered animals go uninspected and for the most part game meats are not inspected.
Biology of Foodborne Parasites
To control and prevent parasitic diseases such toxoplasmosis, trichinellosis, and cysticercosis, care
must be taken to prevent growing animals from exposure to contaminated environments, to ensure that
feed and water are clean and free of parasites, to keep meat scraps and offal away from potential source
animals, and to maintain or develop effective inspection systems.
1.9 Future Directions and Trends
Some estimates indicate that Earth’s human population will reach 9 billion persons by 2050. As part
of the global environmental impact of such a high population, there are concerns for our ability to
provide sufficient quantities of clean water and safe food. Less than 1% of Earth’s freshwater, found in
lakes, rivers, reservoirs, and underground sources shallow enough to be obtained at an affordable cost, is
accessible for direct human use. This is accessible runoff (AC), the water regularly renewed by rain and
snowfall and available on a sustainable basis. AC is unevenly distributed among and within continents
and corresponds poorly to current human population distribution.81 Our present 6 billion population uses
30% of this AC, and billions of people lack basic water services resulting in millions of deaths each year
from water-related diseases.81 By 2025, people on Earth may need to use nearly 70% of AC.81 To meet
this projected demand, global supplies of clean freshwater must be increased. But parasites and other
pathogens from infected humans or animals reach freshwater bodies through unhygienic defecating
habits of humans and the use of human feces for fertilizer and from a wide range of zoonotic reservoirs
such as dogs, cats, cattle, pigs, rodents, birds, and wildlife. Once parasites have reached freshwater,
they can be carried to distant locations or enter into intermediate and definitive hosts such as the fish
or invertebrates we eat, they can attach to irrigated or aquatic plants, or they can contaminate drinking
water. Parasites and other pathogens in water must be better controlled through sanitation, restriction of
access, control of runoff, and more and better water purification methods. Conservation will be essential.
For example, agriculture, the largest user of AC,81 must adopt techniques that make it a more efficient
user. Where water is scarce, unsafe, or of poor quality, basic human needs for water must be viewed as
an international priority.
Fresh produce, especially leafy greens, berries, and fruit juices, are known sources of parasitic infections. From growing through harvesting, washing, packaging, shipping, and marketing to food preparation at home or in commercial venues, many sources of contamination are possible. Removal of zoonotic
reservoir hosts from the proximity to growing areas; easy access to toilet and hand-washing facilities for
growers, pickers, packers, and other food handlers; washing methods designed to remove parasites from
both the produce and from the wash water; and standardized parasite detection methods for foods can all
contribute greatly to improve food safety.
Global production of aquatic products including mollusks, crustaceans, and fish must increase twofold
over the next 25 years to provide the protein needed for a growing population.3 To do this, the aquaculture sector must expand because wild stocks are being overharvested with about 50% of marine fisheries already harvested at maximum yields.6 Although aquaculture can furnish needed protein, provide
employment, and contribute to economic growth, thereby reducing poverty,6 clear guidelines for reducing the introduction of parasites into growing areas can help producers develop improved production
methods, especially in areas where disease is endemic.
According to the World Watch Institute (, pork is the most widely consumed meat in the world, followed by poultry,
beef, and mutton with poultry production the fastest growing meat sector and demand for livestock products expected to nearly double in sub-Saharan Africa and South Asia by 2050. Production ranges from
highly industrialized systems owned by a few multinational companies concentrated in a few countries
to small farms and subsistence producers scattered worldwide. Pressure to increase production in all
these systems can result in serious food safety problems. To prevent and control parasitic diseases such
toxoplasmosis, trichinellosis, and cysticercosis, care must be taken to prevent growing animals from
exposure to contaminated environments, to ensure that feed and water are clean and free of parasites,
to keep meat scraps and offal away from potential source animals, and to maintain or develop effective
inspection systems.
Introduction and Public Health Importance of Foodborne Parasites
1. World Health Organization, Food safety and foodborne illness. Fact sheet no. 237, 2007. http://www.who.
int/mediacentre/factsheets/fs237/en/; Scallon, E. et al., Foodborne illness acquired in the United States—
Unspecified agents. Emerg. Infect. Dis., 17, 16, 2011; Scallon, E. et al., Foodborne illness acquired in the
United States—Major pathogens. Emerg. Infect. Dis., 17, 7, 2011.
2. Keiser, J. and Utzinger, J., Food-borne trematodiases. Clin. Microbiol. Rev., 22, 466, 2009.
3. Keiser, J. and Utzinger, J., Emerging foodborne trematodiasis. Emerg. Infect. Dis., 11, 1507, 2005.
4. World Health Organization, Foodborne trematode infections in Asia. Workshop report RS/2002/
GE/40(VTN), p. 3, 2002.
5. Fürst, T., Keiser J., and Utzinger, J., Global burden of human food-borne trematodiasis: A systematic
review and meta-analysis. Lancet Infect. Dis., 12, 210, 2012.
6. FAO and WHO, Multicriteria-based ranking for risk management of foodborne parasites. Report of a
Joint FAO/WHO Expert Meeting, September 3–7, 2012, FAO Headquarters, Rome, Italy, 2014. http://
7. Batz, M.B., Hoffmann, S., and Morris, J.G. Jr., Ranking the disease burden of 14 pathogens in food
sources in the United States using attribution data from outbreak investigations and expert elicitation.
J. Food Prot., 75, 1278, 2012.
8. Centers for Disease Control.
(Accessed December 2, 2014)
9. Schuster, F.L. and Visvesvara, G.S., Amoebae and ciliated protozoa as causal agents of waterborne zoonotic disease. Vet. Parasitol., 126, 91, 2004.
10. USEPA, Long Term 2 Enhanced Surface Water Treatment Rule (LT2).
11. Baldursson, S. and Karanis, P., Waterborne transmission of protozoan parasites: Review of worldwide
outbreaks—An update 2004–2010. Water Res., 45, 6603, 2011.
12. Tan, K.S., New insights on classification, identification, and clinical relevance of Blastocystis spp. Clin.
Microbiol. Rev., 21, 639, 2008.
13. Leelayoova, S. et al., Evidence of waterborne transmission of Blastocystis hominis. Am. J. Trop. Med.
Hyg., 70, 658, 2004.
14. Eroglu, F. and Koltas, I.S., Evaluation of the transmission mode of B. hominis by using PCR method.
Parasitol. Res., 107, 841, 2010.
15. Lee, I.L. et al., Blastocystis sp.: Waterborne zoonotic organism, a possibility? Parasit. Vectors, 5, 130, 2012.
16. Ortega, Y. and Sanchez, R., Update on Cyclospora cayetanensis, a food-borne and waterborne parasite.
Clin. Microbiol. Rev., 23, 218, 2010.
17. Huang, P. et al., The first reported outbreak of diarrheal illness associated with Cyclospora in the United
States. Ann. Intern. Med., 123, 409, 1995.
18. Marshall, M.M. et al., Waterborne protozoan pathogens. Clin. Microbiol. Rev., 10, 67, 1997.
19. Gonin, P. and Trudel, L., Detection and differentiation of Entamoeba histolytica and Entamoeba dispar
isolates in clinical samples by PCR and enzyme-linked immunosorbent assay. J. Clin. Microbiol., 41,
237, 2003.
20. Pritt, B.S. and Clark, C.G., Amoebiasis. Mayo Clin. Proc., 83, 1154, 2008.
21. Singer, S.M. and Kamda, J., Immune response to Giardia infection: Lessons from animal models.
In: Giardia and Cryptosporidium: From Molecules to Disease, Ortega Pierres, G. et al., eds. CABI,
Cambridge, MA, p. 451, 2009.
22. Yoder, J.S. et al., Giardiasis surveillance—United States, 2009–2010. Morbid. Mortal. Weekly Rep.
Surveill. Summ., 61, 13, 2012.
23. Yoder, J.S., Harral, C., and Beach, M.J., Giardiasis surveillance—United States, 2006–2008. Morbid.
Mortal. Weekly Rep. Surveill. Summ., 59, 15, 2010.
24. Robertson, L.J. and Fayer, R., Chapter 2: Cryptosporidium. In: Foodborne Protozoan Parasites,
Robertson, L.J. and Smith H.V., eds. Nova Science Publishers, Inc., Hauppauge, NY, p. 33, 2012.
25. Mac Kenzie, W.R. et al., A massive outbreak in Milwaukee of cryptosporidium infection transmitted
through the public water supply. N. Engl. J. Med., 331, 161, 1994.
26. Benenson, M.W., Oocyst-transmitted toxoplasmosis associated with ingestion of contaminated water.
N. Engl. J. Med., 307, 666, 1982.
Biology of Foodborne Parasites
27. de Moura, L. et al., Waterborne toxoplasmosis, Brazil, from field to gene. Emerg. Infect. Dis., 307, 666,
28. Waldron, L.S. et al., Molecular epidemiology and spatial distribution of a waterborne cryptosporidiosis
outbreak in Australia. Appl. Environ. Microbiol., 77, 7766, 2011.
29. Weir, M.H. et al., Water reclamation redesign for reducing Cryptosporidium risks at a recreational spray
park using stochastic models. Water Res., 45, 6505, 2011.
30. Hlavsa, M.C., Surveillance for waterborne disease outbreaks and other health events associated with recreational water—United States, 2007–2008. Morbid. Mortal. Weekly Rep. Surveill. Summ., 60, 1, 2011.
31. Cotte, L. et al., Waterborne outbreak of intestinal microsporidiosis in persons with and without human
immunodeficiency virus infection. J. Infect. Dis., 180, 2003, 1999.
32. Yamamoto, N., Risk factors for human alveolar echinococcosis: A case-control study in Hokkaido, Japan.
Ann. Trop. Med. Parasitol., 95, 689, 2001; Painter, J.A., Attribution of foodborne illnesses, hospitalizations, and deaths to food commodities by using outbreak data, United States, 1998–2008. Emerg. Infect.
Dis., 19, 407, 2013.
33. Cook, N. et al., Development of a method to detect Giardia duodenalis cysts on lettuce and the simultaneous analysis of salad products for the presence of Giardia spp. cysts and Cryptosporidium spp. oocysts.
Appl. Environ. Microbiol., 73, 7388, 2007.
34. Centers for Disease Control, Foodborne Outbreak Online Database. US Department of Health and
Human Services, CDC, Atlanta, GA, 2012.
35. Stuart, J.M. et al., Risk factors for sporadic giardiasis: A case-control study in southwestern England.
Emerg. Infect. Dis., 9, 229, 2003.
36. Amahmid, O., Asmama, S., and Bouhoum, K., The effect of waste water reuse in irrigation on the contamination level of food crops by Giardia cysts and Ascaris eggs. Int. J. Food Microbiol., 49, 19, 1999.
37. Smith, H.V. et al., Cryptosporidium and Giardia as foodborne zoonoses. Vet. Parasitol., 149, 29, 2007.
38. Lopez, A.S. et al., Outbreak of cyclosporiasis associated with basil in Missouri in 1999. Clin. Infect. Dis.,
32, 1010, 2001.
39. CDC, Outbreak of cyclosporiasis associated with snow peas—Pennsylvania, 2004. Morbid. Mortal.
Weekly Rep., 53, 676, 2004.
40. Shikanai-Yasuda, M.A. and Carvalho, N.B., Oral transmission of Chagas disease. Clin. Infect. Dis., 54,
845, 2012.
41. ProMed, Archive number: 20120906.1283849.
42. Pereira, K.S. et al., Chagas’ disease as a foodborne illness. J. Food Prot., 72, 441, 2009.
43. Tsai, H.-C. et al., Outbreak of eosinophilic meningitis associated with drinking raw vegetable juice in
southern Taiwan. Am. J. Trop. Med. Hyg., 71, 222, 2004.
44. Ash, L.R., Observations on the role of mollusks and planarians in the transmission of Angiostrongylus
cantonensis infection to man in New Caledonia. Rev. Biol. Trop., 24, 163, 1976.
45. Alatoom, A. et al., Fasciola hepatica in the United States. Lab. Med., 39, 45, 2008.
46. Kern, P. et al., Risk factors for alveolar echinococcosis in humans. Emerg. Infect. Dis., 10, 2088, 2004.
47. Clausen, J.H. et al., Prevention and control of fish-borne zoonotic trematodes in fish nurseries, Vietnam.
Emerg. Infect. Dis., 18, 1438, 2012.
48. FAO FishStat Plus, 2012.
49. Fayer, R. et al., Contamination of Atlantic coast commercial shellfish with Cryptosporidium. Parasitol.
Res., 89, 141, 2003.
50. Lucy, F.E. et al., Biomonitoring of surface and coastal water for Cryptosporidium, Giardia, and humanvirulent microsporidia using molluscan shellfish. Parasitol. Res., 103, 1369, 2008.
51. Freire-Santos, F. et al., Detection of Cryptosporidium oocysts in bivalve molluscs destined for human
consumption. J. Parasitol., 86, 853, 2000.
52. Dorta Contreras, A.J. et al., Eosinophilic meningoencephalitis caused by Angiostrongylus cantonensis
Chen, 1935; immunological study. Rev. Es. Pediatr., 43, 379, 1987.
53. Campbell, B.G. and Little, M.D., The finding of Angiostrongylus cantonensis in rats in New Orleans,
Louisiana, USA. Am. J. Trop. Med. Hyg., 38, 568, 1988.
54. Morera P., Life history and redescription of Angiostrongylus costaricensis Morera and Despedes 1971.
Am. J. Trop. Med. Hyg., 22, 613, 1973.
55. Fried, B. and Abruzzi, A., Food-borne trematode infections of humans in the United States of America.
Parasitol. Res., 106, 1263, 2010.
Introduction and Public Health Importance of Foodborne Parasites
56. CDC, Human paragonimiasis after eating raw or undercooked crayfish—Missouri, July 2006−September
2010. Morbid. Mortal. Weekly Rep., 59, 1573–1576, 2010.
57. Fayer, R. et al., Chapter 42: Waterborne and foodborne parasites. In: Compendium of Methods for the
Microbial Examination of Foods, Vanderzant, C. and Slittstoesser, D.F., eds. American Public Health
Association, Washington, DC, p. 789, 1992.
58. Chai, J.-Y., Murrell, K.D., and Lymbery, A.J., Fish-borne parasitic zoonoses: Status and issues. Int. J.
Parasitol., 35, 1233, 2005.
59. Audicana, M.T. and Kennedy, M.W., Anisakis simplex: From obscure infectious worm to inducer of
immune hypersensitivity. Clin. Microbiol. Rev., 21, 360, 2008.
60. Levsen, A., Lunestad, B.T., and Berland, B., Low detection efficiency of candling as a commonly recommended inspection method for the nematode larvae in the flesh of pelagic fish. J. Food Prot., 68, 828,
61. Ramirez-Avila, L. et al., Eosinophilic meningitis due to Angiostrongylus and Gnathostoma species. Clin.
Infect. Dis., 48, 322, 2009.
62. Behr, M.A. et al., North American liver fluke (Metorchis conjunctus) in a Canadian aboriginal population: A submerging human pathogen? Can. J. Public Health, 89, 258, 1998.
63. Li, M.W. et al., Sparganosis in mainland China. Int. J. Infect. Dis., 15, 154, 2011.
64. Scholz, T. et al., Update on the human broad tapeworm (Genus Diphyllobothrium), including clinical
relevance. Clin. Microbiol. Rev., 22, 146, 2009.
65. FAO, World Livestock 2011—Livestock in Food Security. Food and Agriculture Organization, Rome,
Italy, 2011.
66. Fayer, R., Sarcocystis in human infections. Clin. Microbiol. Rev., 17, 894, 2004.
67. Furtado, J.M. et al., Toxoplasmosis: A global threat. J. Glob. Infect. Dis., 3, 281, 2011.
68. Jones, J.L. and Holland, G.N., Annual burden of ocular toxoplasmosis in the US. Am. J. Trop. Med. Hyg.,
82, 464, 2010.
69. Jones, J.L. et al., Risk factors for Toxoplasma gondii infection in the United States. Clin. Infect. Dis., 49,
878, 2009.
70. Pozio, E., Taxonomy, biology and epidemiology of Trichinella parasites. In: FAO/WHO/OIE Guidelines
for the Surveillance, Management, Prevention and Control of Trichinellosis, Dupouy-Camet, J. and
Murrell, D., eds., p. 1, 2007.
71. Dupouy-Camet, J., Presidential address of ICT12 Conference: “Trichinella and trichinellosis—A never
ending story”. Vet. Parasitol., 159, 194, 2009.
72. Roy, S.L., Lopez, A.S., and Schantz, P.M., Trichinellosis surveillance—United States, 1997–2001.
Morbid. Mortal. Weekly Rep. Surveill. Summ., 52, 1, 2003.
73. Kennedy, E.D. et al., Trichinellosis surveillance—United States, 2002–2007. Morbid. Mortal. Weekly
Rep. Surveill. Summ., 58, 1, 2009.
74. Ramírez-Zamora, A. and Alarcón, T., Management of neurocysticercosis. Neurol. Res., 32, 229, 2010.
75. Eichenberger, R.M., Stephan, R., and Deplazes, P., Increased sensitivity for the diagnosis of Taenia saginata cysticercus infection by additional heart examination compared to the EU-approved routine meat
inspection. Food Control, 22, 989, 2011.
76. USDA, Food Safety Inspection Service Directive, Series 6100.2, p. 17, 2007.
77. Postel, S.L., Daily, G.C., and Ehrlich, P.R., Human appropriation of renewable fresh water. Science, 271,
785, 1996.
78. FDA, Guidance for industry: Guide to minimize food safety hazards of leafy greens; draft g­ uidance.
ProducePlantProducts/ucm064458.htm (December 2, 2014).
79. NOAA, NOAA seafood inspection program. NOAA inspection manual 25, March 2011. http://noaa.ntis.
gov/view.php?pid=NOAA:ocn740058604 (December 2, 2014).
80. USDA, Food Safety Inspection Service directive, Series 6100.2, p.17, 2007.
OPPDE/rdad/FSISDirectives/6100.2.pdf (December 2, 2014).
Molecular Biological Techniques
in Studies of Foodborne Parasites
Una Ryan, Yaoyu Feng, and Lihua Xiao
Introduction..................................................................................................................................... 21
Molecular Detection Methods........................................................................................................ 22
2.2.1 Polymerase Chain Reaction................................................................................................ 22
2.2.2 Quantitative PCR ............................................................................................................... 22
2.2.3 Loop-Mediated Isothermal Amplification and Other Isothermal Amplification Methods.....23
2.2.4 Microfluidic Chips.............................................................................................................. 24
2.2.5 Microarray Detection......................................................................................................... 25
2.3 Molecular Typing and Population Genetics.................................................................................... 25
2.3.1 Multilocus Sequence Typing and Mutation Scanning Typing Methods............................ 25
2.3.2 Population Genetics............................................................................................................ 26
2.4 Phylogenetic Analysis..................................................................................................................... 27
2.4.1 Distance.............................................................................................................................. 27
2.4.2 Parsimony........................................................................................................................... 27
2.4.3 Likelihood Methods........................................................................................................... 28
2.4.4 Rooting Trees..................................................................................................................... 28
2.4.5 Statistical Support for Trees............................................................................................... 28
2.5 Next-Generation Sequencing.......................................................................................................... 29
2.5.1 Pyrosequencing.................................................................................................................. 29
2.5.2 SOLiD (Sequencing by Oligonucleotide Ligation and Detection)..................................... 31
2.5.3 Illumina (Sequencing by Synthesis)................................................................................... 31
2.5.4 Ion Torrent Personal Genome Machine............................................................................. 31
2.5.5 Single-Molecule Real-Time and Nanopore Sequencing.................................................... 32
2.6 Comparative Genomics................................................................................................................... 32
2.7 Genetic Manipulation...................................................................................................................... 33
2.8 Conclusion....................................................................................................................................... 34
Acknowledgment...................................................................................................................................... 34
References................................................................................................................................................. 34
2.1 Introduction
One of the most challenging issues in food safety is the detection of foodborne pathogens. Since the
infectious dose of many pathogens is low, the sensitivity of the diagnostic tool becomes important.
Traditionally, the detection and analysis of foodborne pathogens have relied on light or electron microscopy and culture methods. Conventional techniques such as microscopy lack sensitivity, are labor intensive, and require well-trained microscopists for accurate identification and interpretation, particularly
for pathogens that are morphologically similar or very small in size or present in very low numbers.1–4
Biology of Foodborne Parasites
Special stains are required for the diagnosis of some pathogens. In addition, the diagnostic skills of
microscopists can vary greatly from laboratory to laboratory resulting in some infections being misdiagnosed or missed completely.2,4,5 Furthermore, light microscopy can have a low sensitivity of detection,
which is particularly relevant, considering that, for some pathogens, just a few individuals within a
sample may represent an infectious dose.6,7
In vitro cultivation methods are frequently limited in sensitivity, specificity, or both.4,5,8 Many microorganisms still cannot be cultured, and the isolation of others requires special media and laborious
procedures that cannot be routinely performed in small laboratories.2,8 Culture methods are also very
expensive and time consuming, and in many cases, several days are required before pathogens can be
detected.9,10 In addition, many organisms require strictly controlled conditions during transport of specimens, which also adds to the expense.
Immunoassays have benefits of technical simplicity, rapidity, and cost-effectiveness. Latex particle
agglutination and coagglutination tests, enzyme-linked immunoassays, and direct immunofluorescence
antibody assays have been available for some years. However, immunodiagnostic assays commonly are
hampered by antigenic cross-reactivity (among related or distinct taxa) and low specificity and often do
not allow the distinction among current infection, past infection, and/or exposure.7,11 It has also been
shown recently that some rapid assay kits have low sensitivity in detecting the full range of parasites
within a genus.12 Finally, an important limitation of all conventional diagnostic methods is their inability
to differentiate between different “strains” of the same pathogen.
2.2 Molecular Detection Methods
2.2.1 Polymerase Chain Reaction
The advent of molecular tools, particularly those based on the polymerase chain reaction (PCR), has
provided a major advance for the food industry because of the ability to detect low levels of pathogens
on food. A PCR reaction typically utilizes two oligonucleotide primers, which are hybridized to the
5′ and 3′ ends of the target sequence, and a DNA polymerase, which can extend the annealed primers
by adding on deoxyribonucleoside triphosphates (dNTPs) to generate double-stranded products.13 By
raising and lowering the temperature of the reaction mixture, the two strands of the DNA product are
separated and can serve as templates for the next round of annealing and extension, and the process is
By the early 1990s, the food industry had gained interest in this powerful technique due to its low cost
and ease of use, and numerous primer sets had been developed for the detection of foodborne pathogens.15 Since then, PCR has been used extensively for the detection of foodborne pathogens (see Refs.
[9,16]). For example, PCR was used to detect the presence of Cyclospora DNA in the raspberry filling
of a wedding cake in a Pennsylvania outbreak of cyclosporiasis and for the detection of Cyclospora,
Cryptosporidium, and Giardia in ready-to-eat packaged leafy greens.17,18
Multiplex PCR utilizes more than one set of primers in a reaction and has been used for the simultaneous detection of multiple pathogens in one sample.19,20 There are, however, limitations of the PCR
including inhibitors in foods that can result in false positives. Food-derived PCR inhibitors include Ca2+,
fats, glycogen, and phenolic compounds.21 The presence of proteases in cheese and milk may also inhibit
PCR,22,23 and the detection of Cryptosporidium in water and food samples is often hampered by the
occurrence of organic and inorganic substances that can potentially be PCR inhibitors.24 The use of
bovine serum albumin, different types of DNA polymerase, and different extraction methods has been
used to help overcome this type of inhibition from food.25–27
2.2.2 Quantitative PCR
The invention of quantitative PCR (qPCR) has overcome several limitations of the conventional PCR and
led the way to rapid enumeration of foodborne pathogens.9,10 In qPCR, the amplified product is detected
using fluorescent dyes. These fluorescent dyes are linked to oligonucleotide probes, which bind specifically to the amplified PCR product. Changes in the fluorescence intensities are monitored during the PCR
Molecular Biological Techniques in Studies of Foodborne Parasites
reaction (in “real time”), reflecting the accumulation of the specific PCR product. This not only allows
highly sensitive and specific detection of the target sequences but also makes possible a very accurate
quantitation of the target sequence.10,28 qPCR allows the relative quantification of the amount of DNA
template by using the cycle number at which the fluorescence signal starts to rise above a defined threshold
(termed the Ct value). The Ct value is inversely proportional to the amount of starting DNA template in
the sample, that is, the larger the Ct, the smaller the amount of the starting template. The amount of DNA
template in a sample can be estimated from its Ct by using a standard curve that has been derived from
the Ct’s of appropriate DNA standards containing known amounts of template. The reproducibility of this
measure is normally tested by analyzing replicates of samples and standards in multiple runs.28,29
Nontoxic intercalating dyes (such as SYBR Green, EvaGreen, and SYTO 9) are often used to monitor the accumulation of synthesized amplicon in qPCR. A limitation of intercalating dyes is that they
bind to nonspecific products in an amplicon. Therefore, the specificity of the PCR must be optimized to
minimize nonspecific or erroneous products during the PCR. Increased specificity can be attained using
fluorescent probe–based technologies such as Taqman, molecular beacons, and fluorescent resonance
energy transfer (FRET) probes, which is designed to target the internal sequence of an amplicon and
fluoresces upon hybridization to the target in situ. However, this added specificity incurs increased costs,
as such probes are significantly more expensive than intercalating dyes.30 A downstream melting-curve
analysis with or without probes can be used to increase the specificity of qPCR and for the differentiation of species of genotypes, especially when high-resolution melting-curve (HRM) analysis is used.31,32
A major assumption of qPCR is that all samples will have the same reaction efficiency. This is generally true for high-quality DNA extracts, but as discussed earlier, stool or food samples often yield
DNA of variable quality and may also contain compounds that inhibit PCR or reduce PCR efficiency.
In addition, the well-recognized difficulties in reproducing published tests due to variation in the performance of PCR thermal cyclers33 and the inefficiencies of different DNA polymerases have hampered
implementation of qPCR assays for foodborne parasites in end user laboratories. To overcome these
difficulties, an internal amplification control (IAC) is necessary. An IAC is a nontarget DNA sequence
present in the same sample reaction tube, which is coamplified simultaneously with the target sequence.
The IAC is required to allow differentiation between true negative samples and false negatives due to
PCR inhibition and to allow for any corrections to quantitation estimates to allow for PCR efficiency
differences between the DNA standards and samples. For example, in a PCR without an IAC, a negative
response (no band or signal) can mean that there was no target sequence present in the reaction. But it
could also mean that the reaction was inhibited due to a malfunction of the thermal cycler, incorrect PCR
mixture, poor polymerase activity, and/or the presence of inhibitory substances in the sample matrix.34
Conversely, in a PCR with an IAC, a control signal will always be produced when there is no target
sequence present. When neither IAC signal nor target signal is produced, the PCR has failed. Thus, when
a PCR-based method is used in routine analysis, an IAC, if the concentration is adjusted correctly, will
indicate false-negative results.
To overcome the lack of consensus that exists on how best to perform and interpret qPCR experiments,
the Minimum Information for Publication of Quantitative Real-Time PCR Experiments guidelines have
been developed.35 These guidelines encourage the provision of all relevant experimental conditions and
assay characteristics so that reviewers can assess the validity of the protocols used. Full disclosure of
all reagents, sequences, and analysis methods is necessary to enable other foodborne pathogen detection
laboratories to more easily replicate published protocols.
2.2.3 Loop-Mediated Isothermal Amplification and Other
Isothermal Amplification Methods
Although PCR has been widely used in the detection of foodborne parasites, it requires thermocycling to
separate the two DNA strands, and this characteristic has limited its application in the field. Several isothermal amplification techniques have been developed in the past two decades to overcome the requirement for a thermocycler machine. These techniques include loop-mediated isothermal amplification
(LAMP),36 nucleic acid sequence–based amplification (NASBA),37 rolling circle amplification (RCA),38
and strand displacement amplification.39 NASBA can detect DNA or RNA, and reactions consist of
Biology of Foodborne Parasites
three enzymes including T7 polymerase, dNTPs, and two specific primers and take place at 40°C.37
With RCA, a single forward primer is extended by DNA polymerase along a circular template for many
rounds, displacing upstream sequences and producing a long single-stranded DNA of multiple repeats.
The linear RCA reaction can run for several hours or days, producing millions of copies of the small circle sequence. In exponential RCA, a primer pair is used. The second primer targets the single-stranded
DNA product of the first primer and initiates hyperbranching in the DNA replication, creating as many
as 1012 copies/h.9 In SDA, a primer containing a restriction site is annealed to the DNA template. Next,
amplification primers are annealed to 5′ adjacent sequences (forming a nick) to begin amplification.
Newly synthesized DNA is nicked by the corresponding restriction enzyme, and the polymerase starts
amplification again, displacing the newly synthesized strands. In a single reaction, 109 copies of target
DNA can be produced.39 Of these methods, LAMP is one of the most widely used isothermal amplification assays employed in the detection of foodborne parasites. A major advantage of most isothermal
amplification techniques is that, unlike PCR, this technology is resistant to contamination.
LAMP employs four primers that have a total of six binding sites on the target DNA. It uses a robust
polymerase (BST) to amplify target DNA (or RNA by inclusion of reverse transcriptase) proceeding to
an autocycling strand displacement mechanism, at a constant temperature, producing detectable product
in approximately 1 h.36 It is robust, with reagents stable at ambient temperature for up to 2 weeks40 and
consistently insensitive to extraneous nucleic acids or interference from sample or media components
that are problematic for other nucleic amplification techniques. The procedure is rapid and is able to
amplify from a single copy to 109 in 1 h at constant temperature, typically in the range of 60°C–70°C.36,41
In addition, during LAMP reaction, magnesium pyrophosphate is produced, which increases proportionally the turbidity of the reaction and allows the detection of the product by naked eyes with or without the
addition of fluorescent dye. Detection limits for virus in specimen and media samples have often been
reported as a single copy. LAMP can also be applied to nucleic acid extracts of unpurified samples or
even to samples without nucleic acid extraction, which demonstrates its general insensitivity to extraneous materials other than the target, that is, Toxoplasma oocyst DNA have been detected efficiently in
crude fecal nucleic acid extracts.41 A rapid, sensitive, and specific method for detecting the foodborne
trematode Opisthorchis viverrini from the stool samples using LAMP was developed with results within
40 min, using a heat box or a water bath to maintain the temperature at 65°C.42 LAMP assays have also
been developed for the detection of Cryptosporidium and Giardia in water and feces.43–47 Recently,
LAMP has been shown to be superior to immunofluorescent microscopy and nested PCR detection
of Cryptosporidium in water samples.48 In that study, 25.7% (18/70) of water samples were positive
for Cryptosporidium spp. by immunofluorescent microscopy and 27.1% (19/70) were found positive by
LAMP. Nested PCR products were not generated in any of the investigated water samples. For selected
water pellets spiked with 10 Cryptosporidium oocysts, all 16 samples were positive by LAMP, whereas
only 43.75% (7/16) were positive by nested PCR.48
2.2.4 Microfluidic Chips
Inexpensive, portable, and easy-to-use devices for rapid detection of parasitic pathogens are needed to
ensure safety of food. Microfluidic chips with a multitude of separate reaction wells, each containing
primers for amplification of a specific pathogen, provide a promising platform for multiplexed detection in inexpensive, user-friendly, and compact devices.49 A variety of these chips have been developed
over the years, based on PCR and, more recently, isothermal techniques for DNA/RNA amplification.50–54 More recently, a disposable polymer microfluidic chip for quantitative detection of multiple
pathogens (Salmonella, Campylobacter jejuni, Shigella, and Vibrio cholerae) using isothermal nucleic
acid amplification was developed.55 The chip contains an array of 15 interconnected reaction wells
with dehydrated LAMP primers and required only a single pipetting step for dispensing of sample. For
rapid detection and quantification, LAMP was performed with a highly fluorescent DNA-binding dye
(SYTO 82) and monitored in real time using a low-cost CCD imaging module with an analytical sensitivity of 10–100 gene copies/μL in less than 20 min.55 Multiplexed sensing array for sequence-specific
electrochemical DNA detection of the LAMP products was used in strain discrimination of Salmonella
serovars.52 Due to robustness of LAMP, sample purity is less critical than it is for PCR, which should
Molecular Biological Techniques in Studies of Foodborne Parasites
enable simplified sample preparation procedures that could be more easily performed on-site. Another
option is to integrate sample processing on chip using a variety of microfluidic cell lysis and nucleic
acid purification techniques developed over the years,56 some of which have recently been coupled with
LAMP for detection of infectious agents in clinical samples57–59 and could be applied to foodborne
pathogens in the future.
2.2.5 Microarray Detection
DNA microarray technology has also been explored as a detection tool for foodborne pathogens.
A DNA microarray can contain several thousand surface-immobilized DNA probes on a small glass
slide. Following hybridization of target DNA sequences to probes anchored on the chip surface, fluorescence detection is used to monitor binding events. This approach has been used as a rapid and parallel
high-throughput tool for microbial detection in a single hybridization assay.60 More recently, a random
genomic DNA microarray for the detection and identification of Listeria monocytogenes in milk was
developed with a detection limit of log CFU/mL of L. monocytogenes in milk.61 This type of platform
would be useful for the future development of a DNA microarray that contained random fragments of
genomic DNA of important foodborne pathogens that could be used for rapid and simultaneous detection
of these pathogens in a wide variety of foods.
2.3 Molecular Typing and Population Genetics
The development of molecular methods has provided new tools for enhanced surveillance and outbreak
typing. This has resulted in better implementation of rational infection control programs.62 In outbreak
investigations, a typing method must have the discriminatory power needed to distinguish all epidemiologically unrelated isolates. Ideally, such a method can discriminate very closely related isolates to
reveal person-to-person strain transmission, which is important to develop strategies to prevent further
spread. At the same time, it must be rapid, inexpensive, highly reproducible, and easy to perform and
2.3.1 Multilocus Sequence Typing and Mutation Scanning Typing Methods
PCR coupled with restriction fragment length polymorphisms (PCR–RFLP), length polymorphisms,
sequencing, and/or electrophoretic mutation scanning techniques are widely used for the identification and differentiation of different foodborne species and subtypes within a sample.7 For example,
PCR–RFLP analysis and multilocus typing tools based on microsatellite length polymorphisms have
been developed for Cryptosporidium 65,66 and Clonorchis sinensis, an important foodborne parasite of
humans and animals.7 Although useful, these methods do not detect all of the length and sequence variation within or among amplicons during analysis, and direct DNA sequencing remains the “­gold-standard”
approach for detecting genetic variation or polymorphism. More recently, multilocus sequence typing
(MLST) tools, which involve the PCR amplification and sequencing of multiple diagnostic loci (usually
housekeeping genes), have been developed (e.g., Gatei et al.67), which offer increased resolution because
they are based on sequence analysis of multiple loci.
Mutation scanning techniques such as single-strand conformation polymorphism (SSCP) analysis
have also been developed. Conventional SSCP is performed using nondenaturing gel electrophoresis that
relies on manual scoring of band mobilities against a reference control.68,69 This type of electrophoretic
analysis can, however, be somewhat time consuming to carry out.70 Capillary electrophoresis coupled
with SSCP (CE–SSCP) is a more rapid method and has been used to differentiate between species of
Cryptosporidium but was unable to differentiate between some species within a host group and across
host groups (e.g., C. macropodum and C. canis).71
Mutation scanning methods that rely on the melting characteristics of amplicons and circumvent electrophoretic analysis have also been developed. Melting-curve analysis (an assessment of the dissociation characteristics of double-stranded DNA during heating) has been used to identify and differentiate
Biology of Foodborne Parasites
different foodborne species and subtypes within a sample. The melting profile of a PCR product depends
on its GC content, length, sequence, and heterozygosity,72,73 and the dissociation of double-stranded
DNA is monitored by measuring dye or probe fluorescence as the temperature is slowly and gradually
increased.72 In the case of intercalating dyes such as SYBR Green and SYTO 9, the dissociation of
the DNA strands releases the dye, thus returning it to its nonfluorescent state. Using this process, the
rate of dissociation of the double-stranded DNA template can be measured by a reduction in fluorescence over time, and the temperature that causes the greatest rate of change in fluorescence, corresponding with the period at which double-stranded DNA dissociation is greatest, is inferred to represent the
melting temperature of the amplicon. This approach has been used to detect and differentiate between
Cryptosporidium parvum and C. hominis74 and to differentiate five common Cryptosporidium parasites
that are pathogenic for humans in a single PCR.75
Recent advances in intercalating dyes, improved real-time PCR instrumentation, and analytical
approaches have enabled HRM analysis, allowing the reliable detection of complex melting curves
consisting of multiple peaks representing the gradual dissociation of relatively AT-rich regions
(“lower” melting domains) within the amplicon prior to the ultimate melting and single-nucleotide
polymorphisms,76,77 which can result in slight but measurable changes to the HRM profile by affecting
amplicon dissociation. This approach was used to differentiate between C. hominis, C. parvum, and
C. ­meleagridis.70 Recent studies76,77 have shown that complex melting profiles are reproducible using
different intercalating dyes (although being dependent on dye concentration) and that the display of
multiple peaks is diagnostically informative, providing a unique profile with an increased number of
characters to be used for differentiating species or genotypes. Rasmussen et al. evaluated the computer
program MeltSim ( and reported that while not capable
of exactly predicting DNA dissociation in the presence of an intercalating dye, the program was useful
for identifying regions where melting-curve differences could be exploited for diagnostic melting-curve
assay design.77
2.3.2 Population Genetics
The characterization of the population genetic structure of foodborne parasites is important for understanding the pathobiology of various foodborne pathogens and for tracking sources of infection and
dispersal of pathogens. For example, an analysis of a range of C. parvum and C. hominis isolates from
seven countries at nine loci was used to show that both clonal and panmictic population structures exist
and that the relative contribution of each pattern appeared to vary between different geographic regions
and may be dependent on the prevailing ecological determinants of transmission.74 MLST has been used
to show that only a minority of all Entamoeba histolytica infections progress to the development of clinical symptoms in the host and that population differences exist between the E. histolytica strains isolated
from the asymptomatic and symptomatic individuals.78
A variety of software packages are available for analyzing population structure. DNA Sequence
Polymorphism, DnaSP (, is a software package for the analysis of nucleotide polymorphisms from aligned DNA sequence data.79–89 DnaSP can estimate several measures of
DNA sequence variation within and between populations (in noncoding, synonymous, or nonsynonymous sites or in various sorts of codon positions), as well as linkage disequilibrium, recombination,
gene flow, and gene conversion parameters. Linkage disequilibrium can also be analyzed using LIAN
( Genetic structure (i.e., inferring the presence
of distinct populations, assigning individuals to populations, studying hybrid zones, and identifying
migrants and admixed individuals) can be analyzed using STRUCTURE (http://pritch.bsd.uchicago.
edu/structure.html)82,83 and eBURST ( Levels of taxonomic richness can be
compared using rarefaction using PAST software (, which is a statistical method for estimating the number of taxa expected to be present in a random sample of any size
taken from a given collection. The approach is useful to compare levels of taxonomic richness among
environments, as the taxonomic richness in a sample is dependent on its size and fluctuates stochastically due to sampling variation.85
Molecular Biological Techniques in Studies of Foodborne Parasites
2.4 Phylogenetic Analysis
The accurate identification and characterization of foodborne parasites at different taxonomic levels
have important implications for understanding genetic variation and formulating strategies for disease
control. Phylogenetics (the study of evolutionary relationships among groups of organisms) is based on
the comparison of DNA or protein sequences.86 The results of an analysis can be drawn in a hierarchical
diagram or branching tree, with branches joined by nodes and leading to terminals at the tips of the tree.
The branches in a tree are based on the hypothesized evolutionary relationships (phylogeny) between
organisms. Each member in a branch, also known as a monophyletic group, is assumed to be descended
from a common ancestor.
The first step in constructing a tree is building the dataset, which involves aligning the sequences
of interest with sequences retrieved from public nucleotide databases such as GenBank. A variety of
multiple sequence alignment programs are available; one of the most commonly used is Clustal,87,88
which is available online ( Other sequence alignment
programs include MUSCLE ( Sequence alignments and phylogenetic
analysis can also be conducted using Molecular Evolutionary Genetics Analysis (MEGA) (http://
www.­, which is widely used and available for free for both Windows and Mac OS.
Other downloadable and online phylogenetic analysis tools include Mesquite (http://mesquiteproject., TREECON (http://­, and
( The alignment generated can be edited using a software such as BioEdit (
Once data are aligned, there are many different types of phylogenetic analysis that can be implemented.89 The type of analysis used will be determined by compromise between the length of computational time and the degree of rigor required. The main techniques are distance, parsimony, and
likelihood (including Bayesian analysis).90–92 All three (excluding Bayesian analysis) can be performed
using MEGA listed earlier. A list of and links to 392 phylogeny packages (both free and nonfree) and 54
free web servers are available from
2.4.1 Distance
Distance methods (e.g., distance and minimum evolution) calculate pairwise distances between sequences,
and a clustering algorithm (e.g., neighbor joining [NJ]) is then used to group sequences that are most
­similar. This approach has potential for computational simplicity and, therefore, speed.93 However, distance methods do not allow an analysis of which characters contribute to particular groupings.86,93 As with
other methods, the outcome may depend on the order in which entities are added to the starting tree, but
because only one tree is outputted, it is not possible to examine conflicting tree topologies.
2.4.2 Parsimony
Parsimony and likelihood are more informative because they are character-based methods. They use
the aligned characters (DNA or protein sequences) directly during tree inference, that is, these methods
examine each column of the alignment separately and look for the tree that best accommodates all of
this information.86
Parsimony calculates the number of ancestral mutations between trees and examines the most likely
“trees” based on the smallest number of changes, that is, the “most-parsimonious” tree is the one that
requires the fewest number of evolutionary changes from a common ancestor (e.g., nucleotide ­substitutions)
to explain the sequences. With multiple characters, different groupings may be equally plausible, or equally
parsimonious, and therefore, multiple trees are generated and, if required, combined into a consensus
tree.93 The advantage of this method is that it allows all informative characters to be considered (rather
than being summarized by conversion to a pairwise distance). However, it only works on variant characters, and thus, information from conserved sequence common to all taxa being analyzed is excluded.94
Biology of Foodborne Parasites
2.4.3 Likelihood Methods
In contrast to parsimony, maximum likelihood (ML) analyses compute the probability that a dataset
fits a tree derived from that dataset, given a specified model of sequence evolution. When conducting
ML analysis, the data are compared against a set of models of sequence evolution, and the one that best
describes the observed pattern of sequence variation is chosen.86,93 Two programs in which this can be
performed are Modeltest95 and MrModeltest.96 Alternatively, a user-specified model may be chosen.
This model of sequence evolution is then used in the likelihood analysis. The analysis starts with a
specified tree derived from the input dataset (e.g., an NJ tree) and swaps the branches on the starting tree until the tree with the highest likelihood score (i.e., the best probability of fitting the data) is
gained. This score is a function of both the tree topology and the branch lengths (number of character
state changes). Likelihood analysis, therefore, allows an explicit examination of the assumptions made
about sequence evolution.93 Likelihood methods are the most computationally demanding techniques
for phylogenetic analysis.
Bayesian inference is another likelihood method that is gaining popularity and can be implemented
using the program MrBayes.97,98 Bayesian analysis is also available online at (http://www.­ In Bayesian analysis, a further set of assumptions (termed priors)
are inputted into the original model and the branch swapping algorithms differ.
2.4.4 Rooting Trees
If direct evidence of ancestor–descendant relationships is absent, the direction of change must be inferred
by rooting the trees. The most common method used to root molecular trees is out-group rooting,99 which
compares the character states in the group of interest (the in-group) with those in a group that is closely
related to, but definitely not in, the in-group (the out-group). These differences are used to infer the
direction of character change in the resultant tree. Out-groups are usually selected on the basis of prior
knowledge of the group of interest.
2.4.5 Statistical Support for Trees
One of the most commonly used methods to assess the robustness of the data used for making phylogenetic trees is bootstrapping. This algorithm takes random subsamples of the dataset, builds trees from
each of these, and calculates the frequency with which the various parts of the tree are reproduced in
each of these random subsamples.86 The bootstrap value, therefore, shows the percentage of times that
a clade appears when individual characters in the dataset are randomly removed and replaced with data
from another character from the same dataset, and the analysis is performed again for a specified number
of replications.100 As a general rule, if the bootstrap value for a given interior branch is 95% or higher,
then the topology at that branch is considered “correct” but the values above 60% are considered significant.100,101 Its advantage is that it can be applied to different methods of phylogenetic reconstruction
and that it assigns a probability-like number to every possible partition of the dataset (= branch in the
resulting tree). Its disadvantage is that the support for individual groups decreases as more sequences
are added to the dataset and that it just measures how much support for a partition is in a dataset given
a method of analysis.
There are many examples in the literature where phylogenetic analysis has been used to ­b etter
understand relationships between different strains of foodborne pathogens (e.g., Cao et al.102).
Phylogenetic analysis has also been used to better understanding the public health implications
of Cryptosporidium parasites from animals. For example, phylogeny analysis suggested that
C. ­p arvum found in humans and mammals and C. meleagridis in birds and humans were originally
parasites of rodents and mammals, respectively, but have subsequently expanded their host ranges
to include humans.103 It has also been used to demonstrate that liver fluke (C. sinensis) isolates
from Korea and China were nearly identical over the DNA regions analyzed (18S and ITS rRNA
and cox1).7
Molecular Biological Techniques in Studies of Foodborne Parasites
2.5 Next-Generation Sequencing
First-generation sequencing based on the Sanger chain termination method104 has been one of the main
techniques used to analyze genetic variation in foodborne pathogens. Over the past 30 years, h­ owever,
there has been more than a millionfold improvement in the rate of sequence generation with the
­progression from radio-labeled products using slab gels to fluorescent products and capillary electrophoresis to next-generation sequencing (NGS) technologies.62,105 NGS has transformed genetic investigations by providing a cost-effective way to discover genome-wide variations.62,105 The clear advantage
of NGS over traditional Sanger sequencing is the ability to generate millions of reads in single runs at
comparatively low costs. To construct the complete nucleotide sequence of a genome, multiple short
sequence reads must be assembled based on overlapping regions (de novo assembly) or comparisons
with previously sequenced “reference” genomes (resequencing). Whole-genome sequencing (WGS) is
becoming a powerful and highly attractive tool for epidemiological investigations, and it is highly likely
that in the near future WGS technology for routine clinical use will permit accurate identification and
characterization of foodborne pathogens. The cost of WGS continues to decline, and currently, the
cost of sequencing a small genome can be as little as US$100–150 per isolate.62,106 However, the key
challenge will not be to produce the sequence data but to rapidly compute and interpret the relevant
information from large datasets.
WGS projects will enable the study of the evolution of the pathogenic changes in foodborne pathogens
as they have adapted to new environmental surroundings and will lead to invaluable insights into how the
foodborne pathogens are adapted to their lifestyle.8 NGS technologies will provide additional markers
for epidemiological response and are likely to identify new bacterial, protozoal, or viral pathogens,16,107
particularly as only one-fifth of foodborne illnesses in the United States are of known etiology.108,109
A major limitation to the application of NGS to foodborne parasites and outbreak situations has been
the access to sufficient quantities of material. It may be possible to overcome this limitation, to an extent,
through the use of whole-genome amplification systems,110,111 which allow the synthesis of microgram
quantities of total genomic DNA from minute quantities of starting material (e.g., nanograms). These
systems are particularly attractive for resequencing projects, wherein the overall structure of the genome
is largely established (e.g., for C. parvum, see Abrahamsen et al.112). The advent of “third-generation”
sequencing technologies should overcome many of these difficulties as they are more sensitive and much
less material is required. Many whole-genome sequence assembly methods and software have been
developed since the advent of NGS technologies. For a recent review of de novo genome assembly and
annotation software tools, see Zhang et al.113 and Yandell and Ence.114
2.5.1 Pyrosequencing
Roche 454, which uses pyrosequencing technology, was the first commercially successful next-­generation
system ( The method amplifies DNA inside water droplets in
an oil solution (emulsion PCR), with each droplet containing a single DNA template attached to a single
primer-coated bead that then forms a clonal colony.114 The sequencing machine contains many picolitervolume wells, each containing a single bead and sequencing enzymes. dNTPs are added, and when the
correct base is incorporated, pyrophosphate (PPi) is released, which is then converted into v­ isible light
by luciferase.115 At the same time, the unmatched bases are degraded by apyrase. Then another dNTP is
added into the reaction system, and the pyrosequencing reaction is repeated. In late 2009, Roche combined the GS Junior, a benchtop system, into the 454 sequencing system, which simplified the library
preparation and data processing. The read length is up to 700 bp and takes only ~10 h to complete.57
The disadvantages are the relatively high cost (Table 2.1) and the relatively high error rate in terms of
polybases longer than 6 bp.
Pyrosequencing was recently used for WGS of two L. monocytogenes clinical isolates during a large
foodborne outbreak and provided a more detailed real-time assessment of genetic trait characteristic of
the outbreak strains than could be achieved with routine subtyping methods.116 It has also been used to
87 (read length mode),
99 (accuracy mode)
4–6 h
SMRT Sequencing
(Pacific Bio)
Not yet quantified
15 min–h
Nanopore Sequencing
Up to 5 million
Ion Torrent
1 million
10–24 h
Potential for high
sequence yield,
depending upon
sequencer model and
desired application
Having equipment
that can be very
Up to 3 billion
1–10 days, depending
upon sequencer and
specified read length
Slower than other
Low cost per base,
1.2–1.4 billion
~7 days
More expensive and
impractical for larger
sequencing projects
Long individual reads.
Useful for many
20 min–3 h
Sequencing by
Sequencing by Ligation Chain Termination
Synthesis (Illumina)
(SOLiD Technique)
(Sanger Sequencing)
Real-time sequencing of
Less expensive
Long read size.
single molecules at low
equipment. Fast Fast
cost. Doesn’t damage the
DNA, so in theory the same
molecule can be reanalyzed
Low yield at high accuracy. High error rate
Having runs that
Having equipment that can
are expensive.
be very expensive
Cost per 1 million 2
bases (in US$)
Longest read length.
Fast. No PCR.
Reads per run
Time per run
Read length (bp)
Accuracy (%)
Comparison of NGS Methods
Biology of Foodborne Parasites
Molecular Biological Techniques in Studies of Foodborne Parasites
generate WGS data for Salmonella Newport strains, which enabled a better understanding of the genetic
background of pathogenicity and evolutionary history of S. Newport and also provided additional markers for epidemiological response.101
2.5.2 SOLiD (Sequencing by Oligonucleotide Ligation and Detection)
In this technique, DNA fragments are ligated to adapters, then bound to beads. A water droplet in oil
emulsion contains the amplification reagents with only one fragment bound per bead. DNA fragments
on the beads are amplified by the emulsion PCR. After DNA denaturation, the beads are deposited
onto a glass support surface. In the first step, a primer is hybridized to the adapter. Next, a mixture
of oligonucleotide octamers is hybridized to the DNA fragments and ligation mixture added. In these
octamers, the double of fourth and fifth bases is characterized by one of four fluorescent labels at
the end of the octamer. After the detection of the fluorescence from the label, bases 4 and 5 in the
sequence are thus determined. The ligated octamer oligonucleotides are cleaved off after the fifth
base, removing the fluorescent label, then hybridization and ligation cycles are repeated, this time
determining bases 9 and 10 in the sequence; in the subsequent cycle, bases 14 and 15 are determined;
and so on. The sequencing process is continued in the same way with another primer, shorter by one
base than the previous one, allowing the determination of, in the successive cycles, bases 3 and 4, 8
and 9, and 13 and 14.117,118 The most recent versions have read lengths of 85 bp and data outputs of 30G
per run, but the short read length and the time required (~7 days) are still the major shortcomings119
(Table 2.1).
2.5.3 Illumina (Sequencing by Synthesis)
In this method, DNA molecules and primers are first attached on a slide and amplified with polymerase
so that local clonal DNA fragments, initially coined “clusters,” are formed. To determine the sequence,
four kinds of nucleotides (ddATP, ddGTP, ddCTP, and ddTTP), which contain different cleavable fluorescent dyes, are added, and nonincorporated nucleotides are washed away. A camera takes images of the
fluorescently labeled nucleotides, then the dye along with the terminal 3′ blocker is chemically removed
from the DNA, allowing the next cycle. Unlike pyrosequencing, the DNA chains are extended one nucleotide at a time, and image acquisition can be performed at a delayed moment, allowing for very large
arrays of DNA colonies to be captured by sequential images taken from a single camera. Decoupling
the enzymatic reaction and the image capture allows for optimal throughput and theoretically unlimited
sequencing capacity. This technology is cheap ($0.2 per million bases) and is high throughput, that is, it
can potentially handle thousands of samples simultaneously.119
2.5.4 Ion Torrent Personal Genome Machine
Ion Torrent Personal Genome Machine (PGM) uses standard sequencing chemistry, but with a novel,
semiconductor-based detection system. This method of sequencing is based on the detection of hydrogen ions that are released during the polymerization of DNA, as opposed to the optical methods
used in other sequencing systems. A microwell containing a template DNA strand to be sequenced is
flooded with a single type of nucleotide. When a nucleotide is incorporated into the DNA molecules by
the polymerase, a hydrogen ion is released. By detecting the change in pH, PGM recognizes whether
the nucleotide is added or not. If homopolymer repeats are present in the template sequence, multiple
nucleotides will be incorporated in a single cycle. This leads to a corresponding number of released
hydrogens and a proportionally higher electronic signal, that is, if two nucleotides are added, then
twice the amount of voltage is detected.120 The Ion Torrent PGM was recently used in the characterization of an outbreak of exceptionally virulent Shiga-toxin-producing Escherichia coli O104:H4
centered in Germany, with more than 3000 persons infected.121,122 Studies revealed that the outbreak
strain belonged to an enteroaggregative E. coli lineage that had acquired genes for Shiga toxin 2 and
for antibiotic resistance.122
Biology of Foodborne Parasites
2.5.5 Single-Molecule Real-Time and Nanopore Sequencing
The so-called third-generation sequencing technologies include single-molecule real-time (SMRT)
and nanopore sequencing. Third-generation sequencing has two main characteristics. First, PCR is not
needed before sequencing, which shortens DNA preparation time for sequencing. Second, the signal is
captured in real time, which means that the signal, no matter whether it is fluorescent (SMRT) or electric
current (nanopore), is monitored during the enzymatic reaction of adding nucleotide in the complementary strand, allowing the detection of nucleotide modifications such as methylation because of changes
in nucleotide incorporation kinetics.119
Developed by Pacific Biosciences, SMRT sequencing (PacBio RS) is based on the sequencing by
synthesis approach. The DNA is synthesized in zero-mode waveguides (ZMWs)—small well-like containers with the capturing tools located at the bottom of the well. The sequencing is performed with
use of an unmodified polymerase (attached to the ZMW bottom) and fluorescently labeled nucleotides
flowing freely in the solution. The wells are constructed in a way that only the fluorescence occurring
by the bottom of the well is detected. During the reaction, the enzyme will incorporate the nucleotide
into the complementary strand and cleave off the fluorescent dye previously linked with the nucleotide.
Then the camera inside the machine will capture signal in a movie format in real-time observation.119
Sample preparation is very fast; it takes 4–6 h instead of days. Also, it does not need PCR in the preparation step, which reduces bias and error caused by PCR. Second, the turnover rate is quite fast; runs
are finished within a day. Third, the average read length is >3000 bp, which is longer than that of any
second-generation sequencing technology. Although the throughput of the PacBio RS is lower than a
second-generation sequencer, this technology will be quite useful for foodborne diagnostic laboratories
in the future and has been used in the characterization of an outbreak caused by V. cholerae in Haiti.123
Nanopore sequencing (Oxford Nanopore Technologies®). In this novel method of DNA “strand
sequencing,” DNA is sequenced by threading it through a microscopic pore (nanopore) in a membrane.
An electric current flows through the pore, and DNA passing through the nanopore changes the voltage across the channel. Different DNA bases disrupt the current in different ways, letting the machine
electronically read out the sequence. Each nanopore sequences multiple strands of DNA from solution
in succession, as individual strands are passed through the nanopore by a processive enzyme. DNA and
enzyme are mixed in solution and engaged with the nanopore for sequencing, and once the strand has
been completed, a new strand is loaded into the nanopore for sequencing.124,125
In February 2012, Oxford Nanopore Technologies presented a $900, single-use device (MinION) that
is USB enabled and handheld and can sequence a complete bacteriophage genome.125 A larger desktop
machine (GridION) will handle larger volumes, and clusters of these machines can be used for sequencing whole genomes. The GridION contains 2000 nanopores working to produce 150 mb/h, and the
MinION contains 512 nanopores. Nanopore sensors are purely electrical and can detect DNA in concentrations/volumes that are no greater than what is available from a blood or saliva sample.126 Additionally,
nanopores promise to dramatically increase the read length of sequenced DNA to greater than 10k bases.
Longer read lengths cut the costs and errors associated with shorter read lengths, for which the analysis
must piece together the fragments to reconstruct an estimate of the original sequence.125
The biggest issue is an error rate of 4%, and Oxford Nanopore Technologies is working to reduce the
errors to <1% before its release, which has been delayed to overcome this and other technical issues.
The handheld nanopore device will make it easier to read limited amounts of DNA in field situations
including food processing plants, where inspectors could monitor for contamination of food by various
pathogens, and therefore, as prices reduce even further, it holds great promise for rapid and real-time
monitoring of food.
2.6 Comparative Genomics
In recent years, the whole genomes of many important foodborne parasites have been sequenced to
gain understanding of the basic biology, pathogenesis, and transmission of these pathogens.111,127–131
Recent advances in NGS techniques have made WGS routine for many parasites. This is especially
Molecular Biological Techniques in Studies of Foodborne Parasites
true for the WGS of foodborne helminthes, which although are easy to acquire large quantity of
highly pure DNA from for quenching but have large genomes. The WGS of some noncultural protozoa
such as Cryptosporidium spp. and Cyclospora cayetanensis has been hampered by the difficulties in
acquiring enough purified organisms for NGS analysis. Nevertheless, comparative genomic analysis
is now commonly used in the identification of markers for the development of advanced molecular detection tools and potential targets for the development of new therapeutic and vaccine targets,
even for Cryptosporidium spp.132–134 Significant improvements in our understanding of the transmission and virulence of Toxoplasma gondii came from comparative analysis of different lineages of
The first step of any comparative genomic analysis of WGS data is usually genome assembly:
de novo, the assembly of the millions of NGS reads into larger sequence contigs. In some cases, filtering of reads from contaminants from host and bacterial contamination may be needed prior to sequence
assembly. Various open-source and commercial software can be used, such as Velvet (http://www.ebi., Geneious (, and CLC Genomics Workbench (http://www. The availability of reference genomes of closely related
species often aids the evaluation of the quality of the assembly generated. Most of these software packages also allow the mapping of sequence reads to reference genomes. Reference genomes are also needed
in the ordering and viewing of assembled contigs by chromosome, which can be done using commandline tools such as MUMmer ( or graphical-interface programs such as
Mauve (
Unlike analysis of data from bacteria, there are no established pipelines for comparative genomic
analysis of WGS data from parasites; thus, substantial bioinformatics skills are generally needed,
and some basic knowledge of Unix would help. Many bioinformatics programs are available for multiple alignments of whole genomes, including the aforementioned open-source Mauve and commercial Geneious and CLC Bio packages. Identification and visualization can be done using free Mauve
and Artemis Comparison Tool ( The focus of the
comparative genomic analysis should be on both genome construction and sequence polymorphism.
The former includes gene synteny, reorganizations or rearrangements (inversions, translocations, and
recombination), and major insertions and deletions, whereas the latter includes genome-wide SNP
distribution, identification of highly polymorphic genes or regions, markers or geographic tracking,
and corrections of sequence types with phenotypes (host specificity, virulence, drug resistance, environmental resistance, cyst formation, etc.). As these programs allow the zoom-in to DNA sequence,
reference sequences with annotations of coding regions will facilitate the identification of deletion or
insertion of functionally important genes. Annotation of insertions and higher divergent sequences is
frequently needed to identify biologic significance of genome differences. Phylogenetic (especially
maximum parsimony) analysis of WGS data can be done more easily using genome-wide SNPs than
using whole assemblies.
2.7 Genetic Manipulation
The function of potential targets identified in WGS analysis of foodborne parasites generally requires the
verification of genetic manipulation studies. Significant progresses have been made in functional genetics of some foodborne parasites such as T. gondii, although the genetic manipulation remains very challenging for others such as Cryptosporidium spp.133,137 Various targeted genetic manipulation techniques
have been used in functional genetic studies of foodborne parasites, most of which are revise genetics in
nature, which invokes the alteration or detection of target genes. Homologous recombination is the most
common method used in altering or deleting target genes in protozoan parasites.137 A critical first step
in this genetic manipulation is the introduction of exogenous DNA to the nucleus of the parasite without
affecting the growth and survival of the parasite. Electroporation is commonly used for this.138 An effective marker, such as dihydrofolate reductase–thymidylate synthase for Plasmodium spp. and T. gondii,
is usually needed for the selection of stable transformants without exerting to the laborious and less efficient selection of transfected parasites by negative selection using flow cytometry. This, however, is not
Biology of Foodborne Parasites
practical for some parasites that have no effective therapeutic treatments, such as Cryptosporidium spp.
For some parasites such as T. gondii, due to very high rates of nonhomologous recombination, the integration of insert DNA can be random throughout the genome. Recently, mutant type I and type II strains
of T. gondii have been established by deleting the gene encoding the KU80 protein, thus abolishing the
major pathway of nonhomologous recombination and increase targeted integration through homologous
recombination. The Δku80 strains are now commonly used in reverse genetics of T. gondii.139 Recently,
cosmid libraries with large inserts (~45 kb) have also been used to increase targeted integration by
homologous recombination in T. gondii.140 Knockout mutants can be generated by single crossovers in
the middle of the target gene.
The gene knockout approach cannot be used in studies of proteins necessary for the survival of parasites. For these proteins, their functions are usually studied at the posttranscription level by silencing gene expression, using the RNAi approach. This is especially used in studies of gene functions in
Caenorhabditis elegans and parasitic nematodes Schistosoma spp. and Trypanosoma spp.,141–144 as some
protozoan parasites such as Cryptosporidium spp. and Plasmodium spp. do not have the RNAi machinery such as Dicer and Argonaute, although noncanonical RNAi-related pathways might be present in
Plasmodium.145 A classical RNAi system in theory is existing in T. gondii, but RNAi is rarely used in
functional genetic studies of the parasite.137
2.8 Conclusion
In the era of NGS and WGS, we are experiencing an explosion in the use of advanced molecular biologic
tools in the characterization of foodborne parasites, development of next-generation detection and typing
tools, and studies of functional importance of many so-called hypothetical proteins identified in WGS.
Nevertheless, we are still at the infancy of modern molecular biologic studies of foodborne parasites.
Efforts in this area are hampered by the lagging bioinformatics capacity in analyzing huge volumes of
NGS data, the gene-by-gene approach in reverse genetic analysis of thousands of genes in each organism,
and the lack of effective genetic manipulation and cultivation and animal models for some important protozoan pathogens. As a result, many of the newest molecular biologic tools have not been used widely in
the surveillance and investigation of foodborne illnesses by parasites and control of pathogen contamination in food. They are clearly needed for effective implementation of the Food Safety Modernization Act
in the United States and protection of food supplies elsewhere.
The findings and conclusions in this report are those of the authors and do not necessarily represent the
views of the Centers for Disease Control and Prevention.
1. Lee, S.H., Hwang, S.W., Chai, J.Y., Seo, B.S., 1984. Comparative morphology of eggs of heterophyids
and Clonorchis sinensis causing human infections in Korea. Korean J Parasitol 22, 171–180.
2. Singh, B., 1997. Molecular methods for diagnosis and epidemiological studies for parasitic infections.
Int J Parasitol 27, 1135–1145.
3. Chai, J.Y., 2007. Intestinal flukes. In: Murrell, K.D., Fried, B. (eds.), Food-Borne Parasitic Zoonoses:
Fish and Plant-Borne Parasites, Vol. 11. Springer, New York, pp. 53–115.
4. Josko, D., 2012. Updates in immunoassays: Parasitology. Clin Lab Sci 25, 185–190.
5. van Lieshout, L., Verweij, J.J., 2010. Newer diagnostic approaches to intestinal protozoa. Curr Opin
Infect Dis 23, 488–493.
6. Okhuysen, P.C., Chappell, C.L., Crabb, J.H., Sterling, C.R., DuPont, H.L., 1999. Virulence of three distinct Cryptosporidium parvum isolates for healthy adults. J Infect Dis 180, 1275–1281.
7. Huang, S.Y., Zhao, G.H., Fu, B.Q., Xu, M.J., Wang, C.R., Wu, S.M., Zou, F.C., Zhu, X.Q., 2012.
Genomics and molecular genetics of Clonorchis sinensis: Current status and perspectives. Parasitol Int
61, 71–76.
Molecular Biological Techniques in Studies of Foodborne Parasites
8. Gui, J., Patel, I.R., 2011. Recent advances in molecular technologies and their application in pathogen
detection in foods with particular reference to Yersinia. J Pathog 2011, 310135.
9. Gorski, L., Csordas, A., 2010. Molecular detection: Principles and methods. In: Liu, D. (ed.), Molecular
Detection of Foodborne Pathogens. CRC Press, Boca Raton, FL, pp. 1–15.
10. Hoorfar, J., 2011. Rapid detection, characterization, and enumeration of foodborne pathogens. APMIS
Suppl 133, 1–24.
11. Quílez, J., Sánchez-Acedo, C., Clavel, A., del Cacho, E., López-Bernad, F., 1996. Comparison of an
acid-fast stain and a monoclonal antibody-based immunofluorescence reagent for the detection of
Cryptosporidium oocysts in faecal specimens from cattle and pigs. Vet Parasitol 67, 75–81.
12. Agnamey, P., Sarfati, C., Pinel, C., Rabodoniriina, M., Kapel, N., Dutoit, E., Garnaud, C. et al., 2011.
Evaluation of four commercial rapid immunochromatographic assays for detection of Cryptosporidium
antigens in stool samples: A blind multicenter trial. J Clin Microbiol 49, 1605–1607.
13. Mullis, K.B., Faloona, F.A., 1987. Specific synthesis of DNA in vitro via a polymerase-catalyzed chain
reaction. Methods Enzymol 155, 335–350.
14. Eisenstein, M., 2004. DNA cloning and amplification; Breaking the cycle. Nat Methods 1, 1–2.
15. Harris, L.J., Griffiths, M.W., 1992. The detection of foodborne pathogens by the polymerase chain reaction (PCR). Food Res Int 25, 457–469.
16. Maurer, J.J., 2011. Rapid detection and limitations of molecular techniques. Annu Rev Food Sci Technol
2, 259–279.
17. Ho, A.Y., Lopez, A.S., Eberhart, M.G., Levenson, R., Finkel, B.S., da Silva, A.J., Roberts, J.M., Orlandi,
P.A., Johnson, C.C., Herwaldt, B.L., 2002. Outbreak of cyclosporiasis associated with imported raspberries, Philadelphia, Pennsylvania, 2000. Emerg Infect Dis 8, 783–788.
18. Dixon, B., Parrington, L., Cook, A., Pollari, F., Farber, J., 2013. Detection of Cyclospora, Cryptosporidium,
and Giardia in ready-to-eat packaged leafy greens in Ontario, Canada. J Food Prot 76, 307–313.
19. de Freitas, C.G., Santana, A.P., da Silva, P.H., Gonçalves, V.S., Barros Mde, A., Torres, F.A., Murata, L.S.,
Perecmanis, S., 2010. PCR multiplex for detection of Salmonella Enteritidis, Typhi and Typhimurium and
occurrence in poultry meat. Int J Food Microbiol 139, 15–22.
20. Chen, J., Tang, J., Liu, J., Cai, Z., Bai, X., 2012. Development and evaluation of a multiplex PCR for
simultaneous detection of five foodborne pathogens. J Appl Microbiol 112, 823–830.
21. Wilson, I.G., 1997. Inhibition and facilitation of nucleic acid amplification. Appl Environ Microbiol 63,
22. Rossen, L., Nørskov, P., Holmstrøm, K., Rasmussen, O.F., 1992. Inhibition of PCR by components of food
samples, microbial diagnostic assays and DNA-extraction solutions. Int J Food Microbiol 17, 37–45.
23. Powell, H.A., Gooding, C.M., Garrett, S.D., Lund, B.M., Mckee, R.A., 1994. Proteinase inhibition of the
detection of Listeria monocytogenes in milk using the polymerase chain reaction. Lett Appl Microbiol 18,
24. Skotarczak, B., 2009. Methods for parasitic protozoans detection in the environmental samples. Parasite
16, 1–8.
25. Lantz, P., Hahn-Hägerdal, B., Rådström, P., 1994. Sample preparation methods in PCR-based detection
of food pathogens. Trend Food Sci Technol 5, 384–389.
26. Kreader, C.A., 1996. Relief of amplification inhibition in PCR with bovine serum albumin or T4 gene 32
protein. Appl Environ Microbiol 62, 1102–1106.
27. Abu Al-Soud, W., Râdström, P., 1998. Capacity of nine thermostable DNA polymerases to mediate DNA
amplification in the presence of PCR-inhibiting samples. Appl Environ Microbiol 64, 3748–3753.
28. Higuchi, R., Dollinger, G., Walsh, P.S., Griffith, R., 1992. Simultaneous amplification and detection of
specific DNA sequences. Biotechnology 10, 413–417.
29. Higuchi, R., Fockler, C., Dollinger, G., Watson, R., 1993. Kinetic PCR analysis: Real-time monitoring of
DNA amplification reactions. Biotechnology 11, 1026–1030.
30. Jex, A.R., Smith, H.V., Monis, P.T., Campbell, B.E., Gasser, R.B., 2008. Cryptosporidium-biotechnological
advances in the detection, diagnosis and analysis of genetic variation. Biotechnol Adv 26, 304–317.
31. Hadfield, S.J., Chalmers, R.M., 2012. Detection and characterization of Cryptosporidium cuniculus by
real-time PCR. Parasitol Res 111, 1385–1390.
32. Lalonde, L.F., Reyes, J., Gajadhar, A.A., 2013. Application of a qPCR assay with melting curve analysis
for detection and differentiation of protozoan oocysts in human fecal samples from Dominican Republic.
Am J Trop Med Hyg 89, 892–898.
Biology of Foodborne Parasites
33. Schoder, D., Schmalwieser, A., Schauberger, G., Kuhn, M., Hoorfar, J., Wagner, M., 2003. Physical characteristics of six new thermocyclers. Clin Chem 49, 960–963.
34. Hoorfar, J., Cook, N., Malorny, B., Wagner, M., De Medici, D., Abdulmawjood, A., Fach, P., 2003.
Making internal amplification control mandatory for diagnostic PCR. J Clin Microbiol 41, 5835.
35. Bustin, S.A., Benes, V., Garson, J.A., Hellemans, J., Huggett, J., Kubista, M., Mueller, R. et al., 2009.
The MIQE guidelines: Minimum information for publication of quantitative real-time PCR experiments.
Clin Chem 55, 611–622.
36. Notomi, T., Okayama, H., Masubuchi, H., Yonekawa, T., Watanabe, K., Amino, N., Hase, T., 2000. Loopmediated isothermal amplification of DNA. Nucl Acids Res 28, E63.
37. Compton, J., 1991. Nucleic acid sequence-based amplification. Nature 350, 91–92.
38. Fire, A., Xu, S.Q., 1995. Rolling replication of short DNA circles. Proc Natl Acad Sci USA 92, 4641–4645.
39. Walker, G.T., Little, M.C., Nadeau, J.G., Shank, D.D., 1992. Isothermal in vitro amplification of DNA by
a restriction enzyme/DNA polymerase system. Proc Natl Acad Sci USA 89, 392–396.
40. Thekisoe, O.M., Bazie, R.S., Coronel-Servian, A.M., Sugimoto, C., Kawazu, S., Inoue, N., 2009.
Stability of loop-mediated isothermal amplification (LAMP) reagents and its amplification efficiency on
crude trypanosome DNA templates. J Vet Med Sci 71, 471–475.
41. Karanis, P., Ongerth, J., 2009. LAMP—A powerful and flexible tool for monitoring microbial pathogens.
Trends Parasitol 25, 498–499.
42. Arimatsu, Y., Kaewkes, S., Laha, T., Hong, S.J., Sripa, B., 2012. Rapid detection of Opisthorchis viverrini copro-DNA using loop-mediated isothermal amplification (LAMP). Parasitol Int 61, 178–182.
43. Karanis, P., Thekisoe, O., Kiouptsi, K., Ongerth, J., Igarashi, I., Inoue, N., 2007. Development and preliminary evaluation of a loop-mediated isothermal amplification procedure for sensitive detection of
Cryptosporidium oocysts in fecal and water samples. Appl Environ Microbiol 73, 5660–5662.
44. Bakheit, M.A., Torra, D., Palomino, L.A., Thekisoe, O.M., Mbati, P.A., Ongerth, J., Karanis, P., 2008. Sensitive
and specific detection of Cryptosporidium species in PCR-negative samples by loop-­mediated isothermal
DNA amplification and confirmation of generated LAMP products by sequencing. Vet Parasitol 158, 11–22.
45. Inomata, A., Kishida, N., Momoda, T., Akiba, M., Izumiyama, S., Yagita, K., Endo, T., 2009. Development
and evaluation of a reverse transcription-loop-mediated isothermal amplification assay for rapid and
high-sensitive detection of Cryptosporidium in water samples. Water Sci Technol 60, 2167–2172.
46. Nago, T.T., Tokashiki, Y.T., Kisanuki, K., Nakasone, I., Yamane, N., 2010. Laboratory-based evaluation
of loop-mediated isothermal amplification (LAMP) to detect Cryptosporidium oocyst and Giardia lamblia cyst in stool specimens. Rinsho Byori 58, 765–771.
47. Plutzer, J., Törökné, A., Karanis, P., 2010. Combination of ARAD microfibre filtration and LAMP methodology for simple, rapid and cost-effective detection of human pathogenic Giardia duodenalis and
Cryptosporidium spp. in drinking water. Lett Appl Microbiol 50, 82–88.
48. Koloren, Z., Sotiriadou, I., Karanis, P., 2011. Investigations and comparative detection of Cryptosporidium
species by microscopy, nested PCR and LAMP in water supplies of Ordu, Middle Black Sea, Turkey. Ann
Trop Med Parasitol 105, 607–615.
49. McCalla, S.E., Tripathi, A., 2011. Microfluidic reactors for diagnostics applications. Annu Rev Biomed
Eng 13, 321–343.
50. Asiello, P.J. Baeumner, A.J., 2011. Miniaturized isothermal nucleic acid amplification, a review. Lab
Chip 11(8), 1420–1430.
51. Myers, F.B., Henrikson, R.H., Bone, J., Lee, L.P., 2013. A handheld point-of-care genomic diagnostic
system. PLoS ONE 8, e70266.
52. Patterson, A.S., Heithoff, D.M., Ferguson, B.S., Soh, H.T., Mahan, M.J., Plaxco, K.W., 2013. Microfluidic
chip-based detection and intraspecies strain discrimination of Salmonella serovars derived from whole
blood of septic mice. Appl Environ Microbiol 79, 2302–2311.
53. Reinholt, S.J., Behrent, A., Greene, C., Kalfe, A., Baeumner, A.J., 2013. Isolation and amplification of
mRNA within a simple microfluidic lab on a chip. Anal Chem. 86, 849–856.
54. Zhi, X., Deng, M., Yang, H., Gao, G., Wang, K., Fu, H., Zhang, Y., Chen, D., Cui, D., 2013. A novel HBV
genotypes detecting system combined with microfluidic chip, loop-mediated isothermal amplification
and GMR sensors. Biosens Bioelectron 54C, 372–377.
55. Tourlousse, D.M., Ahmad, F., Stedtfeld, R.D., Seyrig, G., Tiedje, J.M., Hashsham, S.A., 2012. A polymer
microfluidic chip for quantitative detection of multiple water- and foodborne pathogens using real-time
fluorogenic loop-mediated isothermal amplification. Biomed Microdev 14, 769–778.
Molecular Biological Techniques in Studies of Foodborne Parasites
56. Kim, J., Johnson, M., Hill, P., Gale, B.K., 2009. Microfluidic sample preparation: Cell lysis and nucleic
acid purification. Integr Biol (Camb) 1, 574–586.
57. Liu, C., Geva, E., Mauk, M., Qiu, X., Abrams, W.R., Malamud, D., Curtis, K., Owen, S.M., Bau, H.H.,
2011. An isothermal amplification reactor with an integrated isolation membrane for point-of-care detection of infectious diseases. Analyst 136, 2069–2076.
58. Wang, C.H., Lien, K.Y., Wu, J.J., Lee, G.B., 2011. A magnetic bead-based assay for the rapid detection of methicillin-resistant Staphylococcus aureus by using a microfluidic system with integrated loopmediated isothermal amplification. Lab Chip 11, 1521–1531.
59. Wu, Q., Jin, W., Zhou, C., Han, S., Yang, W., Zhu, Q., Jin, Q., Mu, Y., 2011. Integrated glass microdevice
for nucleic acid purification, loop-mediated isothermal amplification, and online detection. Anal Chem
83, 3336–3342.
60. Fukushima, M., Kakinuma, K., Hayashi, H., Nagai, H., Ito, K., Kawaguchi, R., 2003. Detection and
identification of Mycobacterium species isolates by DNA microarray. J Clin Microbiol 41, 2605–2615.
61. Bang, J., Beuchat, L.R., Song, H., Gu, M.B., Chang, H.I., Kim, H.S., Ryu, J.H., 2013. Development of a
random genomic DNA microarray for the detection and identification of Listeria monocytogenes in milk.
Int J Food Microbiol 161, 134–141.
62. Sabat, A.J., Budimir, A., Nashev, D., Sá-Leão, R., van Dijl, J.M., Laurent, F., Grundmann, H., Friedrich,
A.W., 2013. Overview of molecular typing methods for outbreak detection and epidemiological surveillance. Euro Surveill 18, 20380.
63. Struelens, M.J., 1996. Consensus guidelines for appropriate use and evaluation of microbial epidemiologic typing systems. Clin Microbiol Infect 2, 2–11.
64. van Belkum, A., Tassios, P.T., Dijkshoorn, L., Haeggman, S., Cookson, B., Fry, N.K., Fussing, V. et al.,
2007. Guidelines for the validation and application of typing methods for use in bacterial epidemiology.
Clin Microbiol Infect 13(Suppl 3), 1–46.
65. Xiao, L., Morgan, U.M., Limor, J., Escalante, A., Arrowood, M., Shulaw, W., Thompson, R.C., Fayer, R.,
Lal, A.A., 1999. Genetic diversity within Cryptosporidium parvum and related Cryptosporidium species.
Appl Environ Microbiol 65, 3386–3391.
66. Mallon, M., MacLeod, A., Wastling, J., Smith, H., Reilly, B., Tait, A., 2003. Population structures and the
role of genetic exchange in the zoonotic pathogen Cryptosporidium parvum. J Mol Evol 56, 407–417.
67. Gatei, W., Das, P., Dutta, P., Sen, A., Cama, V., Lal, A.A., Xiao, L., 2007. Multilocus sequence typing
and genetic structure of Cryptosporidium hominis from children in Kolkata, India. Infect Genet Evol 7,
68. Jex, A.R., Ryan, U.M., Ng, J., Campbell, B.E., Xiao, L., Stevens, M., Gasser, R.B., 2007a. Specific and
genotypic identification of Cryptosporidium from a broad range of host species by nonisotopic, SSCP
analysis of nuclear ribosomal DNA. Electrophoresis 28, 2818–2825.
69. Jex, A.R., Whipp, M., Campbell, B.E., Caccio, S.M., Stevens, M., Hogg, G., Gasser, R.B., 2007b.
A practical and cost-effective mutation scanning-based approach for investigating genetic variation in
Cryptosporidium. Electrophoresis 28, 3875–3883.
70. Pangasa, A., Jex, A.R., Campbell, B.E., Bott, N.J., Whipp, M., Hogg, G., Stevens, M.A., Gasser, R.B.,
2009. High resolution melting-curve (HRM) analysis for the diagnosis of cryptosporidiosis in humans.
Mol Cell Probes 23, 10–15.
71. Power, M.L., Holley, M., Ryan, U.M., Worden, P., Gillings, M.R., 2011. Identification and differentiation of Cryptosporidium species by capillary electrophoresis single-strand conformation polymorphism.
FEMS Microbiol Lett 314, 34–41.
72. Ririe, K.M., Rasmussen, R.P., Wittwer, C.T., 1997. Product differentiation by analysis of DNA melting
curves during the polymerase chain reaction. Anal Biochem 245, 154–160.
73. Reed, G.H., Kent, J.O., Wittwer, C.T., 2007. High-resolution DNA melting analysis for simple and efficient molecular diagnostics. Pharmacogenomics 8, 597–608.
74. Tanriverdi, S., Tanyeli, A., Başlamişli, F., Köksal, F., Kilinç, Y., Feng, X., Batzer, G., Tzipori, S., Widmer,
G., 2002. Detection and genotyping of oocysts of Cryptosporidium parvum by real-time PCR and melting curve analysis. J Clin Microbiol 40, 3237–3244.
75. Limor, J.R., Lal, A.A., Xiao, L., 2002. Detection and differentiation of Cryptosporidium parasites that are
pathogenic for humans by real-time PCR. J Clin Microbiol 40, 2335–2338.
76. Monis, P.T., Giglio, S., Keegan, A.R., Thompson, R.C., 2005. Emerging technologies for the detection
and genetic characterization of protozoan parasites. Trends Parasitol 21, 340–346.
Biology of Foodborne Parasites
77. Rasmussen, J.P., Saint, C.P., Monis, P.T., 2007. Use of DNA melting simulation software for in silico
diagnostic assay design: Targeting regions with complex melting curves and confirmation by real-time
PCR using intercalating dyes. BMC Bioinform 8, 107.
78. Ali, I.K., Clark, C.G., Petri Jr., W.A., 2008. Molecular epidemiology of amebiasis. Infect Genet Evol 8,
79. Rozas, J., Rozas, R., 1995. DnaSP, DNA sequence polymorphism: An interactive program for estimating
population genetics parameters from DNA sequence data. Comput Appl Biosci 11, 621–625.
80. Rozas, J., 2009. DNA sequence polymorphism analysis using DnaSP. In: Posada, D. (ed.), Bioinformatics
for DNA Sequence Analysis; Methods in Molecular Biology Series, Vol. 537. Humana Press, New York,
pp. 337–350.
81. Haubold, H., Hudson, R.R., 2000. LIAN 3.0: Detecting linkage disequilibrium in multilocus data.
Bioinformatics 16, 847–848.
82. Pritchard, J.K., Stephens, M., Donnelly, P., 2000. Inference of population structure using multilocus
genotype data. Genetics 155, 945–959.
83. Hubisz, M.J., Falush, D., Stephens, M., Pritchard, J.K., 2009. Inferring weak population structure with
the assistance of sample group information. Mol Ecol Resour 9, 1322–1332.
84. Feil, E.J., Li, B.C., Aanensen, D.M., Hanage, W.P., Spratt, B.G., 2004. eBURST: Inferring patterns of
evolutionary descent among clusters of related bacterial genotypes from multilocus sequence typing data.
J Bacteriol 186, 1518–1530.
85. Hughes, J.B., Hellmann, J.J., Ricketts, T.H., Bohannan, B.J., 2001. Counting the uncountable: Statistical
approaches to estimating microbial diversity. Appl Environ Microbiol 67, 4399–4406.
86. Baldauf, S.L., 2003. Phylogeny for the faint of heart: A tutorial. Trend Genet 19, 345–351.
87. Higgins, D.G., Sharp, P.M., 1988. CLUSTAL: A package for performing multiple sequence alignment on
a microcomputer. Gene 73, 237–244.
88. Larkin, M.A., Blackshields, G., Brown, N.P., Chenna, R., McGettigan, P.A., McWilliam, H., Valentin, F.
et al., 2007. Clustal W and Clustal X version 2.0. Bioinformatics 23, 2947–2948.
89. Holder, M., Lewis, P.O., 2003. Phylogeny estimation: Traditional and Bayesian approaches. Nat Rev
Genet 4, 275–284.
90. Page, R.D.M., Holmes, E.C., 1998. Molecular Evolution: A Phylogenetic Approach. Blackwell Science,
Oxford, U.K.
91. Graur, D., Li, W.H., 1999. Fundamentals of Molecular Evolution. Sinauer Associates, Sunderland, MA.
92. Nei, M., Kumar, S., 2000. Molecular Evolution and Phylogenetics. Cambridge University Press,
Cambridge, U.K.
93. Harrison, C.J., Langdale, J.A., 2006. A step by step guide to phylogeny reconstruction. Plant J 45,
94. Stevens, J.R., 2007. Phylogenetic methods for the analysis of parasites and pathogens. In: Tibayrenc, M.
(ed.), Encyclopedia of Infectious Diseases: Modern Approaches. Wiley, Brisbane, Queensland, Australia,
pp. 265–297.
95. Posada, D., Crandall, K.A., 1998. Modeltest: Testing the model of DNA substitution. Bioinformatics 14,
96. Nylander, J.A.A., 2004. MrModeltest v2. Program Distributed by the Author. Uppsala University,
Evolutionary Biology Centre, Uppsala, Sweden.
97. Huelsenbeck, J.P., Ronquist, F., 2001. MRBAYES: Bayesian inference of phylogenetic trees.
Bioinformatics 17, 754–755.
98. Huelsenbeck, J.P., Ronquist, F., Nielsen, R., Bollback, J.P., 2001. Bayesian inference of phylogeny and
its impact on evolutionary biology. Science 294, 2310–2314.
99. Maddison, W.P., Donoghue, M.J., Maddison, D.R., 1984. Outgroup analysis and parsimony. Syst Zool 33,
100. Felsenstein, J., 1985. Confidence-limits on phylogenies—An approach using the bootstrap. Evolution 39,
101. Hillis, D.M., Bull, J.J., 1993. An empirical test of bootstrapping as a method for assessing confidence in
phylogenetic analyses. Syst Biol 42, 182–192.
102. Cao, G., Meng, J., Strain, E., Stones, R., Pettengill, J., Zhao, S., McDermott, P., Brown, E., Allard, M.,
2013. Phylogenetics and differentiation of Salmonella Newport lineages by whole genome sequencing.
PLoS ONE 8(2), e55687.
Molecular Biological Techniques in Studies of Foodborne Parasites
103. Xiao, L., Sulaiman, I.M., Ryan, U.M., Zhou, L., Atwill, E.R., Tischler, M.L., Zhang, X., Fayer, R., Lal,
A.A., 2002. Host adaptation and host-parasite co-evolution in Cryptosporidium: Implications for taxonomy and public health. Int J Parasitol 32, 1773–1785.
104. Sanger, F., Nicklen, S., Coulson, A.R., 1977. DNA sequencing with chain-terminating inhibitors. Proc
Natl Acad Sci USA 74, 5463–5467.
105. Stratton, M.R., Campbell, P.J., Futreal, P.A., 2009. The cancer genome. Nature 458(7239), 719–724.
106. Schloss, J.A., 2008. How to get genomes at one ten-thousandth the cost. Nat Biotechnol 26, 1113–1115.
107. Köser, C.U., Holden, M.T., Ellington, M.J., Cartwright, E.J., Brown, N.M., Ogilvy-Stuart, A.L., Hsu,
L.Y. et al., 2012. Rapid whole-genome sequencing for investigation of a neonatal MRSA outbreak.
N Engl J Med 366, 2267–2275.
108. Scallan, E., Griffin, P.M., Angulo, F.J., Tauxe, R.V., Hoekstra, R.M., 2011a. Foodborne illness acquired
in the United States—Unspecified agents. Emerg Infect Dis 17, 16–22.
109. Scallan, E., Hoekstra, R.M., Angulo, F.J., Tauxe, R.V., Widdowson, M.A., Roy, S.L., Jones, J.L., Griffin,
P.M., 2011b. Foodborne illness acquired in the United States—Major pathogens. Emerg Infect Dis 17,
110. Pinard, R., de Winter, A., Sarkis, G.J., Gerstein, M.B., Tartaro, K.R., Plant, R.N., Egholm, M., Rothberg,
J.M., Leamon, J.H., 2011. Assessment of whole genome amplification-induced bias through highthroughput, massively parallel whole genome sequencing. BMC Genom 7, 216.
111. Sørensen, K.M., Jespersgaard, C., Vuust, J., Hougaard, D., Nørgaard-Pedersen, B., Andersen, P.S., 2007.
Whole genome amplification on DNA from filter paper blood spot samples: An evaluation of selected
systems. Genet Test 11, 65–71.
112. Abrahamsen, M.S., Templeton, T.J., Enomoto, S., Abrahante, J.E., Zhu, G., Lancto, C.A., Deng, M. et al.,
2004. Complete genome sequence of the apicomplexan, Cryptosporidium parvum. Science 304, 441–445.
113. Zhang, W., Chen, J., Yang, Y., Tang, Y., Shang, J., Shen, B., 2011. A practical comparison of de novo
genome assembly software tools for next-generation sequencing technologies. PLoS ONE 6(3), e17915.
114. Yandell, M., Ence, D., 2012. A beginner’s guide to eukaryotic genome annotation. Nat Rev Genet 13,
115. Berka, J., Chen, Y.J., Leamon, J.H., Lefkowitz, S., Lohman, K., Makhijani, V., Rothberg, J., Sarkis, J.,
Srinivasan, M., Weiner, M., 2005. Bead emulsion nucleic acid amplification. U.S. Patent Application,
EP1594980 A2.
116. Foehlich, T., Heindl, D., Roesler, A., 2010. Beads for high-throughput nucleic acid analysis. U.S. Patent,
EP2224015 A1.
117. Gilmour, M.W., Graham, M., Van Domselaar, G., Tyler, S., Kent, H., Trout-Yakel, K.M., Larios, O.,
Allen, V., Lee, B., Nadon, C., 2010. High-throughput genome sequencing of two Listeria monocytogenes
clinical isolates during a large foodborne outbreak. BMC Genom 11, 120.
118. Ansorge, W.J., 2009. Next-generation DNA sequencing techniques. New Biotechnol 25, 195–203.
119. Mardis, E.R., 2008. The impact of next-generation sequencing technology on genetics. Trend Genet 24,
120. Liu, L., Li, Y., Li, S., Hu, N., He, Y., Pong, R., Lin, D., Lu, L., Law, M., 2012. Comparison of nextgeneration sequencing systems. J Biomed Biotechnol 2012, 251364.
121. Rusk, N., 2011. Torrents of sequence. Nat Methods 8, 44.
122. Mellmann, A., Harmsen, D., Cummings, C.A., Zentz, E.B., Leopold, S.R., Rico, A., Prior, K. et al., 2011.
Prospective genomic characterization of the German enterohemorrhagic Escherichia coli O104:H4 outbreak by rapid next generation sequencing technology. PLoS ONE 6, e22751.
123. Rohde, H., Qin, J., Cui, Y., Li, D., Loman, N.J., Hentschke, M., Chen, W. et al., 2011. Open-source
genomic analysis of Shiga-toxin-producing E. coli O104:H4. N Engl J Med 365, 718–724.
124. Chin, C.S., Sorenson, J., Harris, J.B., Robins, W.P., Charles, R.C., Jean-Charles, R.R., Bullard, J. et al.,
2011. The origin of the Haitian cholera outbreak strain. N Engl J Med 364, 33–42.
125. Stoddart, D., Heron, A.J., Mikhailova, E., Maglia, G., Bayley, H., 2009. Single-nucleotide discrimination in immobilized DNA oligonucleotides with a biological nanopore. Proc Natl Acad Sci USA 106,
126. Maitra, R.D., Kim, J., Dunbar, W.B., 2012. Recent advances in nanopore sequencing. Electrophoresis 33,
127. Branton, D., Deamer, D.W., Marziali, A., Bayley, H., Benner, S.A., Butler, T., Di Ventra, M. et al., 2008.
The potential and challenges of nanopore sequencing. Nat Biotechnol 26, 1146–1153.
Biology of Foodborne Parasites
128. Adam, R.D., Dahlstrom, E.W., Martens, C.A., Bruno, D.P., Barbian, K.D., Ricklefs, S.M., Hernandez,
M.M. et al., 2013. Genome sequencing of Giardia lamblia genotypes A2 and B isolates (DH and GS)
and comparative analysis with the genomes of genotypes A1 and E (WB and Pig). Genome Biol Evol 5,
129. Bontell, I.L., Hall, N., Ashelford, K.E., Dubey, J.P., Boyle, J.P., Lindh, J., Smith, J.E., 2009. Whole
genome sequencing of a natural recombinant Toxoplasma gondii strain reveals chromosome sorting and
local allelic variants. Genome Biol 10, R53.
130. Jex, A.R., Liu, S., Li, B., Young, N.D., Hall, R.S., Li, Y., Yang, L. et al., 2011. Ascaris suum draft genome.
Nature 479, 529–533.
131. Mitreva, M., Jasmer, D.P., Zarlenga, D.S., Wang, Z., Abubucker, S., Martin, J., Taylor, C.M. et al., 2011.
The draft genome of the parasitic nematode Trichinella spiralis. Nat Genet 43, 228–235.
132. Tsai, I.J., Zarowiecki, M., Holroyd, N., Garciarrubio, A., Sanchez-Flores, A., Brooks, K.L., Tracey, A.
et al., 2013. The genomes of four tapeworm species reveal adaptations to parasitism. Nature 496, 57–63.
133. Bouzid, M., Hunter, P.R., McDonald, V., Elwin, K., Chalmers, R.M., Tyler, K.M., 2013. A new heterogeneous family of telomerically encoded Cryptosporidium proteins. Evol Appl 6, 207–217.
134. Striepen, B., 2013. Parasitic infections: Time to tackle cryptosporidiosis. Nature 503, 189–191.
135. Li, N., Xiao, L., Alderisio, K., Elwin, K., Cebelinski, E., Chalmers, R., Santin, M. et al., 2014. Subtyping
Cryptosporidium ubiquitum, a zoonotic pathogen emerging in humans. Emerg Infect Dis 20, 217–224.
136. Khan, A., Miller, N., Roos, D.S., Dubey, J.P., Ajzenberg, D., Darde, M.L., Ajioka, J.W., Rosenthal, B.,
Sibley, L.D., 2011. A monomorphic haplotype of chromosome Ia is associated with widespread success
in clonal and nonclonal populations of Toxoplasma gondii. mBio 2, e00228–e00211.
137. Minot, S., Melo, M.B., Li, F., Lu, D., Niedelman, W., Levine, S.S., Saeij, J.P., 2012. Admixture and
recombination among Toxoplasma gondii lineages explain global genome diversity. Proc Natl Acad Sci
USA 109, 13458–13463.
138. Limenitakis, J., Soldati-Favre, D., 2011. Functional genetics in Apicomplexa: Potentials and limits.
FEBS Lett 585, 1579–1588.
139. Rug, M., Maier, A.G., 2013. Transfection of Plasmodium falciparum. Methods Mol Biol 923, 75–98.
140. Rommereim, L.M., Hortua Triana, M.A., Falla, A., Sanders, K.L., Guevara, R.B., Bzik, D.J., Fox, B.A.,
2013. Genetic manipulation in Δku80 strains for functional genomic analysis of Toxoplasma gondii.
J Visual Exp e50598.
141. Brooks, C.F., Johnsen, H., van Dooren, G.G., Muthalagi, M., Lin, S.S., Bohne, W., Fischer, K., Striepen,
B., 2010. The toxoplasma apicoplast phosphate translocator links cytosolic and apicoplast metabolism
and is essential for parasite survival. Cell Host Microbe 7, 62–73.
142. Bhardwaj, R., Krautz-Peterson, G., Skelly, P.J., 2011. Using RNA interference in Schistosoma mansoni.
Methods Mol Biol 764, 223–239.
143. Dalzell, J.J., McVeigh, P., Warnock, N.D., Mitreva, M., Bird, D.M., Abad, P., Fleming, C.C. et al., 2011.
RNAi effector diversity in nematodes. PLoS Neglect Trop Dis 5, e1176.
144. Ericson, M., Janes, M.A., Butter, F., Mann, M., Ullu, E., Tschudi, C., 2014. On the extent and role of the
small proteome in the parasitic eukaryote Trypanosoma brucei. BMC Biol 12, 14.
145. Farias, L.P., Krautz-Peterson, G., Tararam, C.A., Araujo-Montoya, B.O., Fraga, T.R., Rofatto, H.K., Silva
Jr., F.P. et al., 2013. On the three-finger protein domain fold and CD59-like proteins in Schistosoma mansoni. PLoS Neglect Trop Dis 7, e2482.
146. Mueller, A.K., Hammerschmidt-Kamper, C., Kaiser, A., 2014. RNAi in Plasmodium. Curr Pharmaceut
Des 20, 278–283.
Detection of Parasites in Foods
Ynes R. Ortega and Joan M. Shields*
3.1 Introduction......................................................................................................................................41
3.2 Parasites in Food..............................................................................................................................41
3.3 Detection of Parasites in Meats and Seafood................................................................................. 42
3.4 Detection of Parasites in Water....................................................................................................... 42
3.5 Detection of Parasites in Produce................................................................................................... 43
3.6 Future Trends and Conclusion........................................................................................................ 47
In Memoriam............................................................................................................................................ 47
References................................................................................................................................................. 47
3.1 Introduction
Foodborne parasites have been infecting and causing illness to humans since antiquity. As humans
migrated and established in other locations, so did the parasites.1 Calcified helminth eggs were discovered in mummies dating from 1200 BC. The Greeks, particularly Hippocrates (460–375 BC), knew
about worms from fish, and Celsus and Galen (AD 129–200) were familiar with Ascaris, Enterobius,
and Taenia.1 Animals for human consumption can serve as reservoirs, intermediary, or definite hosts of
these parasites. Cooking and processing practices have reduced the risk of acquiring these infections.
Although some foods have not been implicated as vehicles in parasitic foodborne outbreaks, care
and preventive measures should be taken to avoid contamination and illness in consumers. This chapter
describes some of the methods that were and are currently used to detect parasites in three food commodities: meats, water, and fresh produce. Because water is used in food preparation, we are considering
it as an integral part of foods.
3.2 Parasites in Food
The parasites discussed in this chapter are members of the protozoa and the helminths. The first
group includes the amoeba, flagellates, ciliates, and coccidia. The helminths are grouped in three
subgroups: the nematodes, trematodes, and cestodes. Conventional methodology, including microscopy used to detect these parasites in foods, will be described. The targets of detection include
­larvae, eggs, cysts, and oocysts, which are frequently also the infective stages of foodborne parasites.
Detailed descriptions and methods for identification of the most significant parasites of each group
will be discussed in other chapters, and the molecular tools available to detect these parasites will
be covered elsewhere.
In most instances when working with human clinical specimens, parasites are usually present in large
numbers; therefore, detection is less cumbersome than with food and environmental samples. When
* During the preparation of this book chapter, Dr. Joan Shields suddenly passed away on December 12, 2012. This chapter
is dedicated in her memory.
Biology of Foodborne Parasites
analyzing foods, there are major challenges to identify and isolate foodborne parasites. The infectious
dose of parasites tends to be small, and the number of parasites present in contaminated food items is
usually small as well. Since there is no enrichment process to propagate parasites present in foods, procedures for their removal and recovery from foods are extremely important.2 If this removal and recovery
are inadequate, the sensitivity of any detection method will be compromised.
3.3 Detection of Parasites in Meats and Seafood
Contamination of meats can occur when animals ingest the parasites and serve as reservoirs or as
intermediate or paratenic hosts of the parasites. Tissue cysts are produced, or infectious stages of the
parasite are present in the edible tissues. Meats can also be contaminated with fecal material or with
contaminated rinse water during the meat processing phase. In the first instance, where the parasite
is infecting the meat, tissue digestion is usually effective in isolating the tissue cysts (Sarcocystis,
Trichinella, etc.). To sample and identify Trichinella cysts, it is recommended to digest 100 g of muscle
(tongue or diaphragm) free of fascia and fat. A 5 g sample will give a detection limit of one larva per
gram (LPG) of tissue.3 Serological testing has been recommended as well.4 Another option is to feed
the suspected meat to a susceptible host where amplification of the parasite could occur. This is the
case with Toxoplasma where mice or cats have been used successfully for this purpose.5 Water rinses
can be a source of contamination of tissue surfaces. Concentration of the rinses is needed before testing
for the parasites.
Detection of eggs of intestinal flukes, liver flukes, and lung flukes in feces and sputum of food animals
can be accomplished by direct examination of the specimen or by the Kato-Katz method, flotation, or
formalin ethyl acetate sedimentation. Infection in animals can be determined by observation of organs
and isolation of adult flukes within the hepatic and biliary ducts.6 The FLOTAC system has been used
in detection of Fasciola hepatica in sheep. Low numbers of eggs can result in negative results, and
trematode species differentiation by morphology can be difficult.6 Examination of multiple samples is
sometimes required to reach a final diagnosis, particularly in the case of Paragonimus spp. A definite
diagnosis is done by examination of adult worms after treatment.7
Proper inspection of infected carcasses and organs during slaughter will detect hydatid cysts and
cysticerci. If live animals are being examined, heavy infections with cysticerci of Taenia solium can
be determined by examining the tongue of the infected animal. Although this is not the most sensitive
assay, it is certainly easier and is commonly used in the noninspected animal markets of developing
Helminths in fish can be identified based on macro- and microscopical observations. The nematodes
Anisakis and Pseudoterranova larvae usually infect organs of ocean fish. Avoiding certain fish sizes,
species, and harvest areas will reduce the risk of these parasites in cephalopods and fish. If organs are
left in the fish for extended periods, larvae will migrate to the muscle fillet.10 The examination of these
parasites is discussed in more detail in Chapter 1.
Paragonimus spp. can be acquired by ingestion of raw crayfish or crab. The metacercariae (a developmental stage of Paragonimus) can be observed in crabs or crayfish. However, this method is not practical
in the seafood industry. Proper cooking is, therefore, vital to inactivating these parasites.11 To find the
metacercaria (the infective stage of this parasite) in the crustaceans, the caparace (hard shell) is removed
to gain access to the gills. The gills are removed, pressed between two glass plates, and examined using a
microscope. The hepatopancreas tissue can also be removed, homogenized, and filtered, and a sediment
is allowed to form. The sediment is then examined by microscopy.12
3.4 Detection of Parasites in Water
If water is contaminated with parasites, then any foods, including vegetables and meats, that are in
contact with it can have cross contamination. This is particularly significant when foods are consumed
raw or minimally processed. Irrigation water has been tested at the farm level using the EPA method
Detection of Parasites in Foods
1623 as well as by flocculation. Ultrafiltration was used to detect protozoan parasites in various types of
water, particularly those for human consumption.13 Many of the procedures can probably be modified
and adapted in the analysis of concentrates or washes from food.
The use of biosensors to detect parasites in water has also been reported. Giardia cysts can be immobilized with monoclonal antibodies and detected using a piezoelectric-excited millimeter-sized cantilever
(PEMC) biosensor. One to ten cyst/mL can be detected in various water matrices (buffer, tap, and river
water) without a preconcentration step.14 In another study, an electrochemical biosensor based on a polymer substrate was developed with the benefits of being cost effective and disposability of the materials.
Amplified hsp70 mRNA from Cryptosporidium parvum using NASBA has been used in a sandwich
hybridization assay with capture probe-coated superparamagnetic beads and reporter probe-tagged liposomes.15 The liposomes entrapped potassium ferro-/ferrihexacyanide to enable amperometric quantification of the amplicon. The investigators reported detection of mRNA from as few as one oocyst using this
poly methyl methacrylate (PMMA) biosensor.
The EPA 1623 method and the integrated cell culture–PCR (CC-PCR) technique have been compared
for efficacy in detecting Cryptosporidium in six watersheds.16 The success in detection was 58.5% for the
CC-PCR technique and 72% for the EPA 1623 method.
Novel filtration methodologies have also been described for parasite recoveries. The counterflow
microrefinery (CFMR) system consists of multiple counterflow concentration units. By using various units, concentration from 10 and 100 L can be reduced to mL with no recirculation processing.17
Recoveries of 81.3% Cryptosporidium oocysts and 86.2% Giardia cysts from water containing as few
as 0.5–100 organisms/L were achieved. Water turbidities and sample volumes were not significant problems in using this method. A field-deployable device using microbead immunoagglutination combined
with Mie scatter detection in a microfluidic device has been developed.18 This method was able to capture
small fragments of the Cryptosporidium cell wall (COWP), detecting fewer than one oocyst, even when
15 µL of the water sample was tested using the microfluidic biosensor.
In addition to the conventional microscopic and immunoassays, novel molecular methods have been
used not only to detect but also to determine the viability of the oocysts. Tissue culture and mouse
neonate models have been used to address oocyst viability, but they are cumbersome, expensive, and
time consuming. Molecular methods have been developed to overcome these challenges. For example,
propidium monoazide–qPCR targeting hsp70 gene was compared to reverse transcription (RT)-qPCR
heat-induced hsp70 mRNA in water containing aged oocysts before and after disinfection using ammonia or hydrogen peroxide.19 PMA–DNA assay was not as sensitive as the mRNA assay in detecting
viability alterations.
Pathogen contamination of vegetables can occur if contaminated soil is in contact with the edible parts. Methods to detect oocysts in soil samples have also been addressed. A qPCR protocol
for detection and quantification of C. parvum in natural soil matrices and soil–water extracts was
developed. Minimum detection limits of 0.667 and 6.67 for fresh and aged C. parvum oocysts were
achieved, respectively, and the percent recovery of Cryptosporidium oocysts ranged from 4.3% to
107.8% for three types of soil samples.20
3.5 Detection of Parasites in Produce
Contamination of produce can occur when irrigation or fumigation water containing parasites,
infected food handlers and processors, and contaminated soil and reservoirs (insects/wildlife/­
domestic animals) come into contact with and possibly adhere to the edible part of the produce. This
contamination can occur at the various stages of the food chain, from the farm to the table. Humans
acquire infection when contaminated, minimally processed produce is ingested. This is particularly important when protozoa can survive in water on the surface of fresh produce. Produce contaminated with Cyclospora have caused outbreaks in North America in 1996–1997, 2013, and 2014.
Produce associated with these outbreaks include fresh berries, cilantro, and salad mix.21 Moreover,
the presence of Cyclospora, Cryptosporidium, and Giardia in precut salads and leafy greens raises
concern that parasites are perhaps more frequently present in produce than previously considered.
Biology of Foodborne Parasites
100 um
30 um
FIGURE 3.1 Scanning electron microscopy of a raspberry. Trichomes are covering the surface and observed at low
(a) and high magnifications (b).
Between April 2009 and March 2010, 544 retail samples (salad blends and leafy greens) were collected and tested for the presence of parasites. Most of these products were grown in the United States.
Parasites were detected using PCR, sequencing, and immunofluorescence microscopy. Cyclospora
spp. was identified in 1.7%, Cryptosporidium spp. in 5.9%, and Giardia in 1.8% of the samples. It is
important to determine the sources of contamination of these vegetables and if these parasites are
viable and infectious.22
It is assumed that in most instances parasites are located on the surface of the produce; however, there
is some evidence that small parasites (e.g., Cryptosporidium) can enter the stoma, making even more
difficult their removal from fresh produce.23 In other instances, the surface of edible plant parts has a
complex surface topography, for example, the presence of trichomes in raspberries (Figure 3.1), which
adds a physical barrier for parasite removal. In addition, the hydrophobicity and charge of the edible
parts of the plants will limit the access of water-based wash solutions, thereby making the removal of the
parasites more difficult.23
Since parasites cannot be enriched, removal from foods and produce (vegetables, fruits, and herbs)
is critical to achieving detection. Identification of the parasite using two methods, direct observation (macro- and microscopic analyses) and molecular tools, is usually recommended. Adherence is
another factor to consider. Some parasites (Cyclospora) adhere more strongly than others (Giardia) to
the vegetable surfaces (Ortega, unpublished observations). Several wash solutions and concentration
methods have been described for protozoa and eggs of helminths. In the BAM FDA method, detection of Cyclospora and Cryptosporidium on produce is achieved by washing (by gentle agitation) with
distilled water.24 A filter bag facilitates the separation of the plant material from the rinses. Other
studies suggest the use of elution buffers or the use of detergents to remove parasites from the surface
of vegetables.25 In other situations, the vegetable or fruit is homogenized and the parasites are then
concentrated (Table 3.1).
Various studies have addressed the role of fresh produce in transmitting of parasites. In Spain, the
occurrence of Giardia and Cryptosporidium on salad products and in irrigation waters was determined.
Water (100 mL) from irrigation canals samples was concentrated using immmunomagnetic separation
(IMS) and FA staining. Four water samples were taken weekly 1 month before harvesting the vegetables. The waters tested were positive for both Cryptosporidium and Giardia. The salad products
were washed with 1 M glycine (pH 5.5) and stomached for 30 s, followed by IMS and FA cyst/oocysts
staining. Cryptosporidium was detected in 12 of 19 samples and Giardia was detected in 10 of 19
samples. Recoveries of Texas Red–stained Cryptosporidium and Giardia, used as internal controls, were
24.5%±3.5% for Cryptosporidium and 16.7%±8.1% for Giardia.26
30–100 g lettuce, bean sprouts
50 g bean sprouts and salad mix,
100 g strawberries, Chinese leaves,
lettuce (iceberg and green lollo),
red cabbage, carrots
30 g lettuce, 60 g raspberries
14–150 g of various salads, lettuce,
chilies, carrots, vegetable stir fry,
parsley, spring onions, watercress,
spinach, pak choi, baby sweet corn,
Lettuce for experimental inoculation.
Cabbage, celery, cilantro, green
onions, leeks, lettuce, parsley, black
mint, yerba buena, and ground
green chili for market sampling
50 g bean sprouts, 100 g iceberg
lettuce, mushrooms, raspberries
Sample Volume
Food Matrix
200 mL extractant (0.1 M HEPES, 1 M sodium
bicarbonate, pH 6.0, 1 M glycine, pH 5.5, 1 M bicine,
pH 5.6, 1% lauryl sulfate, EB [EPA method 1623]), 0.1 M
tricine, pH 5.4, PBS, pH 7.2. Removal by (a) 1 min
stomaching, (b) pulsification for 1 min, (c) orbital shaking
(80 rpm) for 1 min, and (d) spiral rolled for 1 min.
200 mL 1 M glycine, pH 5.5. Stomaching for 30 s,
concentrated by centrifugation at 2500 g × 10 min.
150 mL water and 50 mL elution buffer (EPA method
1623), repeated twice. Drum rotated for 1 or 5 min,
200 mL water and 50 mL elution buffer. 1 min rotating
drum at 80 rpm for 1 min, sonicated for 3 min.
Concentrated by centrifugation at 2500 g × 10 min. 10 mL
was used for IMS using WGA-coated paramagnetic beads,
10 mL for direct observation after centrifugation.
200 mL water and 50 mL elution buffer. 1 min rotating
drum at 80 rpm for 1 min, sonicated for 3 min.
Concentrated by centrifugation at 2500 g × 10 min. 10 mL
was used for IMS using WGA-coated paramagnetic beads,
10 mL for direct observation after centrifugation.
50 mL distilled water, handshaking.
Wash Solution/Elution, Concentration
Methods for the Detection of Protozoan Parasites in Fresh Produce
IMS (Dynbeads GC combo IMS test kit
[Dynal, U.K.]), Giardia-a-Glo™ and
Crypt-a-glo™ FITC (Waterborne™,
Inc., New Orleans, LA)
IMS Crypto-Scan (ImmuCell Co.,
Portland, ME), and IFA Crypt-a-glo™
FITC (Waterborne™, Inc., New
Orleans, LA)
IMS (Dynabeads antiCryptosporidium, Dynal A.S.);
Dynabeads GC Combo, Dynal, A.S.;
Crypto-Scan, ImmuCell Co.,
Portland, ME. IFA (AquaGlo G/C
direct, Waterborne)
IMS-Dynal A.S. Oslo, Norway, and
bright-field microscopy for Al
Epifluorescence microscopy
Acid-fast stain, epifluorescence, IFA
MERIFLUOR Cryptosporidium/
Giardia (Meridian Bioscience, Inc.,
Cincinnati, OH)
Giardia, Ascaris
(Continued )
Detection of Parasites in Foods
1 kg strawberries, 0.5 kg carrots,
20 radishes, 1 lettuce
50 g raspberries, 25 g leafy greens,
25 g spring mix lettuce
25 g leafy greens
25 g basil
10–25 g leafy produce, 50 g berries
10–25 g leafy produce, 50 g berries
100 g raspberries
Orange/ apple juice, cider, milk
(milk products), 25 mL, pH 8.0
Orange apple juice, cider, milk
(milk products), 10 mL
Sample Volume
Food Matrix
Microscopy, epifluorescence
PCR amplification, quantitative
real-time PCR
Microscopy, Qiagen Dneasy Blood and
Tissue Kit, PCR–RFLP, and
200 mL of water, 3% levulinic acid/3% SDS, 1 M glycine, FastDNA® Spin for Soil Kit (MP
PBS, 0.01% Alconox, or 100 mL of 1% HCl/6.4% pepsin. Biomedicals, Irvine, CA), PCR
amplification, microscopy, sequencing
100 mL dH2O. Centrifuge to 45 mL. 2.5 mL of 20% Celite in FTA filter and FTA purification buffer,
NET BSA, 1% polyvinyl polypyrrolidone (PVPP), decant
PCR, epifluorescence
celite in pass through a grade 4 filter by vacuum. Wash
filtrate with NET.
100 mL dH2O. Centrifugation, 0.5 mL pellet in 10 mL.
IMS—Dynabead anti-Cryptosporidium
Kit, IFA Hydrofluor Combo Detection
Kit for Giardia cysts and
Cryptosporidium oocysts (Ensys, Inc.,
Research Triangle Park, NC)
100 mL distilled water. Agitated for 30 min, centrifuged.
Poly-Prep chromatography column
with tightly packed glass wool.
Filtrate applied to FTA filter
(Ftizco, Inc., Maple Plain, MN); PCR
Add same volume of NET buffer. 2.5 mL of 20% Celite in FTA filter and FTA purification buffer,
NET BSA, 1% PVPP, decant celite in pass through a
PCR, epifluorescence
grade 4 filter by vacuum. Wash filtrate with NET.
Centrifugation, 0.5 mL pellet in 10 mL.
IMS—Dynabead anti-Cryptosporidium
Kit, IFA Hydrofluor Combo Detection
Kit for Giardia cysts and
Cryptosporidium oocysts (Ensys, Inc.,
Research Triangle Park, NC)
2 L distilled water and 20 mL Tween 20. 20 mL 1 M
CaCl2 solution, 20 mL NaHCO3, 1 M NaOH to final pH
10. Left overnight, pellet suspended in 200 mL 10%
NH3O3S, 0.01% T-80 washed, pelleted, and reconstituted.
Deionized water, 1 M glycine, pH 5.5, 0.1% Alconox.
Rocked for 15 min twice, centrifuged.
PBS, 0.01% Tween 80, pH 7.4.
Wash Solution/Elution, Concentration
Methods for the Detection of Protozoan Parasites in Fresh Produce
TABLE 3.1 (Continued )
Biology of Foodborne Parasites
Detection of Parasites in Foods
3.6 Future Trends and Conclusion
Molecular methods are increasingly utilized to detect parasites in food and water because of their sensitivity and feasibility for future molecular typing. Real-time and reverse transcriptase PCR techniques
have been developed for most parasites. Still, the gold standard for identification of parasites is the identification of the parasitic stages in food matrices. Another challenge for parasite detection is to determine
the public health significance of the pathogen detection; that is, whether parasites isolated from foods
are viable and if they are infectious to humans. The use of animals to propagate some parasites isolated
from foods can be achieved; however, this approach is time consuming, costly, and cumbersome. As for
detection during inspection, conventional methods are still being used, particularly with parasitic helminths. The development of diagnostic tests that are fast, cost efficient, and without a doubt confirmatory
is needed not only by the regulatory agencies but also by the food industry.
In Memoriam
Dr. Joan Shields (1964–2012)
Dr. Shields was a parasitologist in the U.S. Food and Drug Administration’s Center for Food Safety and
Applied Nutrition. Her studies at the FDA led to improved detection methods for foodborne parasites such
as Cyclospora. Prior to joining the FDA, she was with the Centers for Disease Control and Prevention,
where she studied Cryptosporidium in recreational water. She passed away on December 12, 2012.
1. Cox, F.E., History of human parasitology, Clin. Microbiol. Rev., 15, 595, 2002.
2. Ortega, Y.R. and Sanchez, R., Update on Cyclospora cayetanensis, a food-borne and waterborne parasite,
Clin. Microbiol. Rev., 23, 218, 2010.
3. Forbes, L.B. et al., Complete validation of a unique digestion assay to detect Trichinella larvae in horse
meat demonstrates the reliability of this assay for meeting food safety and trade requirements, J. Food
Prot., 71, 558, 2008.
4. Gamble, H.R. et al., International Commission on Trichinellosis: Recommendations on the use of serological tests for the detection of Trichinella infection in animals and man, Parasite, 11, 3, 2004.
5. Dubey, J.P. et al., High prevalence and genotypes of Toxoplasma gondii isolated from organic pigs in
northern USA, Vet. Parasitol., 188, 14, 2012.
6. Keiser, J. and Utzinger, J., Food-borne trematodiasis: Current chemotherapy and advances with artemisinins and synthetic trioxolanes, Trends Parasitol., 23, 555, 2007.
Biology of Foodborne Parasites
7. Sayasone, S. et al., Epidemiology of Opisthorchis viverrini in a rural district of Southern Lao PDR,
Trans. Roy. Soc. Trop. Med. Hyg., 101, 40, 2007.
8. The Cysticercosis Working Group in Peru, The marketing of cysticercotic pigs in the Sierra of Peru, Bull.
World Health Organ., 71, 223, 1993.
9. Gonzalez, A.E. et al., Prevalence and comparison of serologic assays, necropsy, and tongue examination
for the diagnosis of porcine cysticercosis in Peru, Am. J. Trop. Med. Hyg., 43, 194, 1990.
10. Adams, A.M., Murrell, K.D., and Cross, J.H., Parasites of fish and risks to public health, Rev. Sci. Tech.,
16, 652, 1997.
11. CDC, Paragonimiasis. http//, accessed November 26, 2014.
12. Nawa, Y. et al., Paragonimus westermani and Paragonimus species. In Encyclopedia of Food Safety,
Motarjemi, Y., Moy, G., and Todd, E. (eds.). Academic Press, San Diego, CA, pp. 179–187, 2014.
13. Hill, V.R. et al., Comparison of hollow-fiber ultrafiltration to the USEPA VIRADEL technique and
USEPA method 1623, J. Environ. Qual., 38, 822, 2009.
14. Xu, S. and Mutharasan, R., Rapid and sensitive detection of Giardia lamblia using a piezoelectric cantilever biosensor in finished and source waters, Environ. Sci. Technol., 44, 1736, 2010.
15. Nugen, S.R. et al., PMMA biosensor for nucleic acids with integrated mixer and electrochemical detection, Biosens. Bioelectron., 24, 2428, 2009.
16. LeChevallier, M.W. et al., Comparison of method 1623 and cell culture-PCR for detection of
Cryptosporidium spp. in source waters, Appl. Environ. Microbiol., 69, 971, 2003.
17. Pires, N.M., Recovery of Cryptosporidium and Giardia organisms from surface water by counter-flow
refining microfiltration, Environ. Technol., 34, 2541, 2013.
18. Angus, S.V., Kwon, H.J., and Yoon, J.Y., Field-deployable and near-real-time optical microfluidic biosensors for single-oocyst-level detection of Cryptosporidium parvum from field water samples, J. Environ.
Monit., 14, 3295, 2012.
19. Liang, Z. and Keeley, A., Comparison of propidium monoazide-quantitative PCR and reverse transcription quantitative PCR for viability detection of fresh Cryptosporidium oocysts following disinfection and
after long-term storage in water samples, Water Res., 46, 5941, 2012.
20. Koken, E. et al., Quantification of Cryptosporidium parvum in natural soil matrices and soil solutions
using qPCR, J. Microbiol. Methods, 92, 135, 2013.
21. Cyclosporiasis outbreak investigations—United States, 2013.­cyclosporiasis/​
outbreaks/investigation-2013.html, accessed February 21, 2014.
22. Dixon, B. et al., Detection of Cyclospora, Cryptosporidium, and Giardia in ready-to-eat packaged leafy
greens in Ontario, Canada, J. Food Prot., 76, 307, 2013.
23. Macarisin, D., Bauchan, G., and Fayer, R., Spinacia oleracea L. leaf stomata harboring Cryptosporidium
parvum oocysts: A potential threat to food safety, Appl. Environ. Microbiol., 76, 555, 2010.
24. Orlandi, P.A. et al., Detection of Cyclospora and Cryptosporidium, Bacteriological Analytical Manual., accessed November
26, 2014.
25. Shields, J.M., Lee, M.M., and Murphy, H.R., Use of a common laboratory glassware detergent improves
recovery of Cryptosporidium parvum and Cyclospora cayetanensis from lettuce, herbs and raspberries,
Int. J. Food Microbiol., 153, 123, 2012.
26. Amorós, I., Alonso, J.L., and Cuesta, G., Cryptosporidium oocysts and giardia cysts on salad products
irrigated with contaminated water, J. Food Prot., 73, 1138, 2010.
27. Ortega, Y.R. et al., Isolation of Cryptosporidium parvum and Cyclospora cayetanensis from vegetables
collected in markets of an endemic region in Peru, Am. J. Trop. Med. Hyg., 57, 683, 1997.
28. Robertson, L.J., Gjerde, B., and Campbell, A.T., Isolation of Cyclospora oocysts from fruits and vegetables using lectin-coated paramagnetic beads, J. Food Prot., 63, 1410, 2000.
29. Robertson, L.J. and Gjerde, B., Factors affecting recovery efficiency in isolation of Cryptosporidium
oocysts and Giardia cysts from vegetables for standard method development, J. Food Prot., 64, 1799,
30. Robertson, L.J. and Gjerde, B., Isolation and enumeration of Giardia cysts, Cryptosporidium oocysts,
and Ascaris eggs from fruits and vegetables, J. Food Prot., 63, 775, 2000.
31. Cook, N. et al., Towards standard methods for the detection of Cryptosporidium parvum on lettuce and
raspberries. Part 1: Development and optimization of methods, Int. J. Food Microbiol., 109, 215, 2006.
Detection of Parasites in Foods
32. Cook, N. et al., Development of a method for detection of Giardia duodenalis cysts on lettuce and for
simultaneous analysis of salad products for the presence of Giardia cysts and Cryptosporidium oocysts,
Appl. Environ. Microbiol., 73, 7388, 2007.
33. Lass, A. et al., The first detection of Toxoplasma gondii DNA in environmental fruits and vegetables
samples, Eur. J. Clin. Microbiol. Infect. Dis., 31, 1101, 2012.
34. Chandra, V., Torres, M., and Ortega, Y.R., Efficacy of wash solutions in recovering Cyclospora cayetanensis, Cryptosporidium parvum, and Toxoplasma gondii from basil, J. Food Prot., 77, 1348, 2014.
35. Orlandi, P.A. and Lampel, K.A., Extraction-free, filter-based template preparation for rapid and sensitive
PCR detection of pathogenic parasitic protozoa, J. Clin. Microbiol., 38, 2271, 2000.
Section II
Important Foodborne Protists
Christen Rune Stensvold
Introduction..................................................................................................................................... 53
Morphology and Classification....................................................................................................... 54
4.2.1 Life Cycle and Morphology............................................................................................... 54
4.2.2 Blastocystis Taxonomy....................................................................................................... 55
4.2.3 Blastocystis Subtypes......................................................................................................... 55
4.3 Biology, Genetics, and Genomics................................................................................................... 56
4.3.1 Genome of Mitochondrion-Like Organelle........................................................................ 59
4.3.2 Nuclear Genome................................................................................................................. 59
4.4 Diagnosis and Typing...................................................................................................................... 60
4.4.1 In Vitro Isolation of Blastocystis.........................................................................................61
4.4.2 Subtyping.............................................................................................................................61
4.4.3Impact of Sampling and Sample Processing on Blastocystis Detection and Subtyping....... 62
4.5 Epidemiology and Molecular Epidemiology.................................................................................. 63
4.5.1 Molecular Epidemiology of Blastocystis........................................................................... 63
4.5.2 Linking Molecular Epidemiology to Clinical Presentation............................................... 64
4.6 Pathogenesis and Clinical Features................................................................................................ 65
4.6.1 Blastocystis and Intestinal Eukaryotic Microbiota............................................................ 67
4.6.2 Blastocystis and Irritable Bowel Syndrome....................................................................... 68
4.6.3 Blastocystis Animal Models.............................................................................................. 68
4.6.4 In Silico Prediction of Blastocystis Pathogenicity by Genome Analysis.......................... 69
4.7 Treatment and Prevention............................................................................................................... 69
4.8 Concluding Remarks and Future Directions.................................................................................. 70
Acknowledgment...................................................................................................................................... 70
References................................................................................................................................................. 70
4.1 Introduction
Blastocystis1,2 is a common, strictly anaerobic, unicellular intestinal parasitic protist of humans and a
vast variety of nonhuman hosts.3,4 Despite the fact that it is one of the most common microeukaryotes to
colonize the human intestine and its clinical significance is largely unknown, it remains a relatively little
studied parasite. This may be due in part to a variety of unrelated predicaments, including (1) the inconspicuousness and elusiveness of the parasite, making diagnosis difficult; (2) the fact that it is extremely
difficult to isolate in axenic culture; (3) the resilience of colonization, which may be chronic and difficult
to eradicate; (4) the inclination of the parasite to present itself along with other intestinal microeukaryotes
such as Dientamoeba, making it difficult to know whether potential symptoms are due to Blastocystis
or any other organism; and (5) the fact that Blastocystis genetically is not even remotely related to other
microeukaryotes colonizing or infecting the human intestine, such as protozoa and yeasts.
Biology of Foodborne Parasites
Blastocystis has been known for slightly more than 100 years. By mid-February 2013, a search on
Blastocystis among articles indexed at PubMed retrieved 988 entries, 518 (52.4%) of which had been
published within the 10 most recent years, showing that interest in the parasite has increased significantly
lately. During the past few years, the focus has been centered partly on the development of improved
methods for diagnosis and molecular characterization, and substantial advances in Blastocystis research
have been made; these were recently reviewed3 and mainly include (1) standardization of nomenclature,
(2) molecular epidemiological surveys of Blastocystis in different human cohorts across the globe and in
wild and captive animals, (3) molecular characterization of Blastocystis isolates by highly discriminatory molecular tools, and (4) sequencing and annotation of mitochondrial and nuclear genomes. Aspects
of Blastocystis evolution and biology have been subject to scrutiny, and new hypotheses regarding the
parasite’s role in health and disease are being developed.
This chapter aims to provide an account of many of the knowns and unknowns of Blastocystis with a
major emphasis on the diagnosis and the molecular epidemiology of the parasite; this will be topped off
by addressing perspectives of future research in the light of advances in relevant technologies.
4.2 Morphology and Classification
4.2.1 Life Cycle and Morphology
In humans, Blastocystis is a parasite of the colon and it is shed, possibly intermittently,5 in stool by
colonized individuals. While it has been widely accepted that Blastocystis is transmitted by fecal–oral
transmission (in contaminated food or water or by direct contact), the particular stages in its life cycle
have been the subject of a contentious debate; for a description of the many morphotypes reported to
date, please refer to Refs. [6–8].
It has been maintained that avacuolar, multivacuolar, and amoeboid stages may all represent vegetative in vivo stages.6,9 Other stages such as the vacuolar and granular stages may to a varying extent represent artifacts influenced by changes in physiochemical conditions such as exposure to oxygen.10 On the
other hand, it is clear that the vacuolar stage, which is the stage mostly seen in well-maintained cultures,
can often also be detected by direct microscopy of freshly passed stool and by permanent staining of
fixed fecal smears, and the subject of the in vivo stage(s) remains controversial.3,7
The stage responsible for transmission is most probably the cyst. Cysts may be shed by at least
20%–30% Blastocystis carriers,11,12 but intermittent shedding is likely and so point prevalence data probably underestimate the proportion of cyst shedders. They are round to ovoid and are within the range
of 3.5–12.65 µm12–15 with no particular morphological hallmarks apart from the presence or absence of
a surface coat (fibrillar layer), which may account for the reported differences in cyst size. Cysts may
develop into vacuolar stages in vitro within 24 h.16,17 While vacuolar stages detected in xenic in vitro
culture (XIVC) may have developed from the cysts, noncyst stages present in feces may also (re)develop
into vacuolar stages in XIVC, since cysts are not a consistent finding in fecal samples positive by XIVC.12
The vacuolar stage is the one typically found in primary or well-maintained older cultures. It may also be
encountered in direct or stained fixed fecal smears or in fecal concentrates where sample processing may distort morphology significantly. Iodine staining of this stage reveals a large vacuole inside the cell, which may
be involved in storage of carbohydrates and/or lipids. The size of the vacuolar stage may very well depend on
the health of the parasite; in well-maintained cultures (i.e., in cultures where the medium is replaced twice
a week and most of the cells discarded, a process known as subculturing), usually small and very homogeneous cells are seen (5–10 µm); in neglected cultures where subculturing is not performed, the cells appear
to gain dramatically in size, and cells maybe become as large as 200 µm (giant cells). Under such conditions,
the cell population becomes very heterogeneous with cells differing by size and appearance. For instance, a
stage referred to generally as the “granular stage” is common in neglected cultures; they are generally larger
than the vacuolar stage and may represent degrading cells originating from the vacuolar stage.10,18
The “amoeboid stage” has infrequently been reported.19 However, since there is currently no standardized description of the different morphological stages, it is generally difficult to interpret published data on
morphotypes. Authors who are not familiar with the variety of morphotypes described in the literature may
think of an “amoeboid stage” as a general term for a vegetative, noncyst stage, that is, similar to the term
“trophozoite,” whereas other authors may distinguish between amoeboid, avacuolar, multivacuolar, vacuolar, and granular stages. For example, a paper by Zuel-Fakkar et al.19 describes finding the amoeboid form
in almost two-thirds of patients with Blastocystis and urticaria; however, the authors do not report which
stage(s) was/were found in the remaining one-third; this makes interpretation of the finding impossible.
4.2.2 Blastocystis Taxonomy
In 1996, Silberman et al.20 took to generating complete Blastocystis small subunit (SSU) rDNA sequences
(1.8 kbp) and submitted them to phylogenetic analysis. The parasite, which had been considered anything
from yeast to protozoon, was eventually shown to belong to the heterogeneous group of organisms called
stramenopiles, which are uni- and multicellular protists, consisting especially of algae (from diatoms to
kelp) but also oomycota (water molds), including one of the most significant plant pathogens, namely,
Phytophthora, the cause of potato blight. The phylogenetic position of Blastocystis was later confirmed
by phylogenetic analysis of other genes, including concatenated mitochondrial nad genes.21–23
So far, a few genera from this vast group of organisms have been identified as being parasitic, anaerobic endobionts, some examples being Opalina and Proteromonas, which include intestinal parasites of
anurans and lizards, respectively. Proteromonas has been found in the hindgut of lizards and, interestingly, Proteromonas DNA was recently demonstrated in blood and tissue samples from lizards.24 One
other member of the stramenopiles is known to be able to parasitize on humans, namely, Pythium insidiosum; human pythiosis is characterized primarily by granulomatous disease with a variety of clinical
From an evolutionary biology standpoint, it is highly interesting that a stramenopile has so successfully adapted to a parasitic life style, capable of protractedly colonizing a plethora of very diverse host
species including members of primates, other mammals, birds, reptiles, amphibians, and arthropods and
thereby evading innate and adaptive immune defenses from such a diverse range of hosts.
4.2.3 Blastocystis Subtypes
The remarkable intrageneric diversity of Blastocystis was called into attention in 1997. 25,26 Based
on SSU rDNA analysis, it appears that humans are natural hosts of nine distinct, morphologically indistinguishable lineages of Blastocystis (Table 4.1), currently named subtypes (STs),
Blastocystis STs in Humans by Geographic Region
64 (24.1)
1 (14.3)
17 (6.4)
120 (45.1)
6 (85.7)
6 (2.3)
41 (15.4) 18 (6.8)
38 (44.2)
31 (36.0)
17 (19.8)
281 (32.4) 118 (13.6) 368 (42.4)
9 (1.0)
7 (0.8) 15 (1.7)
70 (8.1)
279 (40.3) 26 (3.8)
11 (1.6)
11 (1.6)
2 (0.3)
18 (1.6)
1 (1.0)
8 (0.7)
2 (1.9)
1 (0.1)
882 (27.8) 343 (10.8) 1399 (44.1) 318 (10.0) 9 (0.3) 89 (2.8)
118 (3.7) 10 (0.3) 3 (0.1)
338 (48.8)
186 (16.2) 146 (12.7) 506 (44.0)
33 (32.0) 5 (4.9)
44 (42.7)
25 (3.6)
278 (24.2) 2 (0.2) 4 (0.3)
14 (13.6) 0
4 (3.9)
Source: Adapted from Alfellani, M.A. et al., Acta Trop., 126, 11, 2013.
Notes: Percentages indicated in parentheses; mixed/untypable cases not included.
Biology of Foodborne Parasites
which may in fact be considered nine distinct species. 3,27 All these lineages except ST9 have also
been found in nonhuman hosts.
Many species names can be encountered in the Blastocystis literature. Isolates from reptiles have been
named Blastocystis cycluri, Blastocystis lapemi, Blastocystis geochelone, and Blastocystis pythoni, and
from the rodent group, there is Blastocystis ratti. Until 2007, Blastocystis from human, many mammalian, and avian hosts were named Blastocystis hominis. Some reptilian and amphibian isolates seem
to fall within the range of genetic variation covered by the mammalian and avian lineages, and if the
reptilian and amphibian isolates are distinct species, the nine lineages of Blastocystis found in humans
cannot all be B. hominis.27 Likewise, B. ratti (which has later been identified as belonging to the ST4 lineage) has been found in both humans and nonhuman hosts, and the use of this name renders B. hominis
a paraphyletic species due to the aforementioned same reasons. Therefore, a consensus terminology was
reached in 2007.27 It was decided that separate lineage by named “subtypes” and that Blastocystis from
humans, other mammals, and birds be named Blastocystis sp. followed by an ST number, depending on
the lineage in question. At that time, nine STs (ST1–ST9) were included in the ST system.
While the number of STs colonizing humans remains stable, many new STs have been discovered in
nonhuman hosts over the past few years, primarily by amplification of Blastocystis-specific DNA directly
from fecal DNA template by PCR and sequencing; to date, 17 STs have been identified (Figure 4.1).
In 2009, Stensvold et al. reported on ST10 in cattle and two ring-tailed lemurs.28 This ST has more
recently been shown to be one of the predominant STs in artiodactyls (Table 4.2)28–31; like some other
STs, it comprises at least two major genotypes that differ genetically by 1% (in the bar-code region, see
Section 4.4).
Later, Parkar et al.32 identified three new STs: ST11 was found in elephants, ST12 in giraffes and
kangaroos, and ST13 in quokkas (the marsupial Setonix brachyurus); ST13 was later found in wild
Tanzanian colobus monkeys, although originally described as ST5,3,33 and also in a mouse deer.29
ST14 was reported in cattle by Fayer et al.31 and later in cattle, sheep, camels, and a mouflon,29 and
recently, three novel STs were reported in nonhuman hominoids (gibbon and chimpanzee), a lemur
and a camel (ST15), kangaroos (ST16), and a gundi (ST17).29 Interestingly, ST15 and ST17 are phylogenetically closer related to reptilian or arthropod isolates than to STs usually found in mammalian
hosts (Figure 4.129).
Hence, the genetic universe and host range of Blastocystis appears to be expanding before us, and
further sampling from various types of hosts will assist in identifying transmission patterns and host
specificity and moreover test the validity of the ST nomenclature. At some point, this may enable us
to generate species names based on observations on host specificity and genetic diversity. Fortunately,
the past few years have seen a tremendous rise in the interest in Blastocystis molecular epidemiology,
and ST surveys in humans and animals are currently emerging from many regions across the globe
(see Section 4.5).
4.3 Biology, Genetics, and Genomics
The in vivo activity of Blastocystis and the consequences thereof remain an enigma. A surface coat
containing various types of carbohydrates may be used for trapping and degrading bacteria.8 Clusters
of bacteria have been observed inside Blastocystis by light microscopy, although it may be difficult to
identify whether this is due to phagocytosis or bacterial invasion (unpublished observations). It has been
suggested that Blastocystis control bacterial numbers by inhibition or predation.34 However, to the author’s
knowledge, there is little evidence of phagocytosis in Blastocystis,35 and the parasite may possibly acquire
its nutrients from the host environment mainly or exclusively by pinocytosis36 or transmembrane transporters.37 Related species such as Opalina (colonizing anurans) and Proteromonas (colonizing lizards) are
also parasites of the large intestine in hosts where nutrient absorption occurs in the small intestine, and so
Blastocystis may not compete with its host in terms of nutrient acquisition.
Blastocystis sp. Nandll (U51151; Human)
Blastocystis sp. (AB107968; Vervet monkey)
Blastocystis sp. (AF439782; Human)
Blastocystis sp. (AB107961; Pig)
Blastocystis sp. (AB107969; Pig-tailed macaque)
Blastocystis sp. (EU445487; Pig)
59/0.77/* 100/1.0/100
Blastocystis sp. (AB070987; Human)
Blastocystis sp. (AY618265; Human)
Blastocystis sp. GEPA2 (EF209020; Leopard tortoise)
Blastocystis sp. Bg1 (GU256922; Elephant)
Blastocystis sp. (AY266469; Toad)
Blastocystis sp. (AB107966; Cattle)
Blastocystis sp. (AB107964; Pig)
Blastocystis sp. (EF468654; Human)
Blastocystis sp. Mouflon
Blastocystis sp. Cow1 (Cattle)
Blastocystis sp. 930296 (GU256902; Giraffe)
Blastocystis sp. 417 (Mousedeer)
Blastocystis sp. 49 (GU256934; Quokka)
Blastocystis sp. (AB091237; Human)
Blastocystis sp. (EU445485; Chicken)
Blastocystis sp. (AY135411; Turkey)
Blastocystis sp. (Human_cid_paper)
Blastocystis sp. (AF408426; Human)
Blastocystis sp. (AY590107; Human)
Blastocystis sp. (AF408427; Human)
Blastocystis sp. (AB070991; Human)
Blastocystis sp. (AB107963; Pig)
Blastocystis sp. (AB107965; Cattle)
Blastocystis sp. (HQ909889; Human)
Blastocystis sp. (AB070992; Human)
Blastocystis sp. (HQ909890; Baboon)
Blastocystis sp. CA6 (Camel)
Blastocystis sp. (AB107970; Lemur)
Blastocystis sp. (AB107971; Pheasant)
Blastocystis sp. (U51152; Guinea pig)
Blastocystis sp. (AY135412; Duck)
Blastocystis sp. MKJ04-10 (EU427512; Red kangaroo)
Blastocystis sp. MKJ04-30 (EU427514; Red kangaroo)
Blastocystis cycluri (AY266474; Rhino iguana)
99/1.0/99 Blastocystis sp. (EU082109; Human)
Blastocystis sp. (DQ366343; Human)
Blastocystis sp. (AB071000; Rat)
Blastocystis sp. (JN6826513; Human)
Blastocystis sp. AFJ96-T8 (AY266467; Leopard frog)
Blastocystis lapemi (AY266471; Sea snake)
Blastocystis pythoni (AY266472; Reticulated python)
Blastocystis sp. R44 (AY266475; Rhino lguana)
Blastocystis sp. GERA3a (EF209017; Radiated tortoise)
Blastocystis sp. GECA2 (EF209018; Red-footed tortoise)
Blastocystis sp. KINIX2 (EF209019; Bell’s hinge-backed tortoise)
Blastocystis sp Gundi
Blastocystis sp. C4 (DQ186645; Cockroach)
Blastocystis sp. C4 (DQ186646; Cockroach)
Blastocystis sp. GERA3b (EF209016; Radiated tortoise)
Blastocystis sp. MA7 (Gibbon)
Blastocystis geocheloni (AY266473; Red-footed tortoise)
Blastocystis sp. (AFJ96-U12; Bullfrog)
Blastocystis sp. AFJ96-H12 (AY266470; Toad)
FIGURE 4.1 Phylogenetic relationship among 17 Blastocystis STs based on neighbor-joining analysis of the SSU rDNA
(30.2) (10.9) (37.7) (8.1)
(10.0) (0.3)
(1.1) (0.1) (1.6)
(0.3) (0.1)
(29.4) (50.0) (2.9)
Sources: Modified after papers by Alfellani, M.A. et al., Acta Trop., 126, 11, 2013; Alfellani, M.A. et al., Parasitology, 140(8), 966, 2013.
Note: Mixed/untypable cases not included.
Homo sapiens 882
Perissodactyls 1
Distribution of Blastocystis STs by Host Group
Biology of Foodborne Parasites
Of the more exciting scientific advances in recent years has been the realization that many types of
commensal microorganisms, including microeukaryotes, are not simple “passengers” in the bodies of
humans and animals but instead have key roles in physiological processes, including immune responses
and metabolism, as well as in disease. Some ciliates, for instance, are commonly found to interfere with the
metabolism of carbohydrates and short-chain fatty acids (SCFAs) in ruminants (please consult references
by Pandey et al.34 and Kittelmann et al.39); we do yet not know whether and how Blastocystis is capable of
influencing host metabolism and how and its immune-stimulating role remains scarcely investigated.
4.3.1 Genome of Mitochondrion-Like Organelle
Blastocystis, which is strictly anaerobic, has organelles that resemble mitochondria, the so-called
­mitochondrion-like organelles (MLOs). These are possibly characterized best as “obligately anaerobic
mitochondria” and may represent an intermediate stage along the degenerative evolutionary pathway
from mitochondrion to derived organelles such as hydrogenosomes or mitosomes.23 Initially, analysis
of more than 12,000 EST sequences revealed that metabolic pathways in Blastocystis are characterized
by the remarkable absence of cytochrome and ATPase subunits, hence confirming previous observations40,41; Blastocystis is the only stramenopile studied so far that lacks all the genes for cytochromes,
and it is hypothesized that selection for maintaining cytochrome complex genes was lost in Blastocystis
during a facultatively anaerobic stage.23 While many of the pathways present are characteristic of classical mitochondria, these organisms also share some features reminiscent of hydrogenosome-containing
species, such as a gene-encoding [FeFe] hydrogenase. The so-called iron/sulfur (Fe/S) cluster biosynthetic machinery is a key pathway in bacteria and eukaryotes and necessary for various essential cell
functions, among which are electron transport, enzymatic catalyses, and regulation of gene expression.
Blastocystis is in possession of genes associated with the synthesis of Fe/S clusters that may have been
acquired by an ancestor of Blastocystis by lateral gene transfer from a methanoarchaeon, some of which
are common inhabitants of the human intestine.42
MLO genome sequences are available for ST1, ST2, ST3, ST4, and ST7 (unpublished observations).23,41,43,44 Genomes are circular, and sizes range from 27.7 to 29.3 kbp with identical gene content
and arrangement existing across the different STs. For comparison, mitochondrial genomes of other
stramenopiles are, not surprisingly, larger and range from 31.6 to 58.5 kbp.45
MLO genome phylogeny reflects the phylogeny obtained by SSU rDNA sequence analysis in terms
of phylogenetic position and relationship between STs23 (unpublished observations). The remarkable
genetic diversity in Blastocystis existing at SSU rRNA gene level is possibly even more pronounced at
the MLO genome level; for instance, the overall nucleotide identity between the strains DMP/02-328
(ST4) and NandII (ST1) MLO genomes is 75.4% or 71.6% and 64.3% for nucleotides and amino acids,
respectively, when only protein-coding genes are considered.23
The MLO genome is A+T rich (80%) and contains 45 genes, of which 27 are protein coding; the
remainder represent structural RNAs such as tRNAs (n = 16) and small and large subunit ribosomal
RNAs. Protein-coding genes include nad genes (n = 10), ribosomal protein-coding genes (n = 13), and
four open reading frames for which gene orthologs are yet to be identified. The reduced set of tRNA
genes implies that tRNAs for some codons must be nuclear encoded and imported from the cytoplasm.
The in silico MLO proteome identifies 365 proteins, 299 of which are mitochondrial-import proteins; 41
of these proteins may be involved in hitherto undiscovered metabolic processes38; an in silico reconstruction of metabolic pathways of Blastocystis MLO was recently published.38
Genetic loci in MLO genomes from ST3 and ST4 have recently been used for multilocus sequence
typing (MLST)44 (see Section 4.5).
4.3.2 Nuclear Genome
While the Blastocystis MLO genome holds information on key components of some essential pathways
such as electron transport and translation, studies of the nuclear genome may provide some insight into,
for instance, potential effector proteins. The only complete genome published to date is the one from
the “B” strain (ST7) sequenced by traditional Sanger sequencing and published by Denoeud et al.38
Biology of Foodborne Parasites
The genome comprises 18.8 Mb and is possibly the smallest stramenopile genome sequenced to date;
genomes of other stramenopiles range from 27.4 to 240.0 Mb.3 Fifteen chromosomes have been identified by pulsed-field gel electrophoresis, and the genome comprises 6020 genes corresponding to 42% of
the genome38; the ploidy of Blastocystis remains unknown. Little overall synteny between Blastocystis
and other stramenopile species is seen, which in part may be due to frequent gene rearrangements
(recombination). Analysis of the genome of ST7 revealed the presence of 133 genes that might be of bacterial or archaeal origin and that have been acquired by horizontal gene transfer (HGT), and moreover,
there are indications of HGT from pathogenic/anaerobic eukaryotes; enzymes involved in fermentation
are closely related to homologues found in, for example, Trichomonas vaginalis and Entamoeba histolytica.38 Other genes possibly acquired by HGT include some coding for hydrolases and proteins related
to tissue adherence.
The in silico secretome comprises 307 proteins, 75 of which are proteins that in some other organism
have been related to pathogenicity, including proteases, hexose digestion enzymes, lectins, glycosyltransferases, and protease inhibitors.38
It is hardly surprising, given the extent of genetic divergence, that little synteny exists between
Blastocystis and other stramenopiles, and therefore, studies of genomes of other STs should merit high
priority. Levels of synteny/recombination should be compared and so should panels of genes coding
for effector proteins. Analysis of the genome of Proteromonas lacertae, one of the closest relatives of
Blastocystis identified to date, is ongoing and will also assist in the description of the evolutionary history of stramenopiles adapting to the life of anaerobic endobionts and potentially reveal differences/
similarities in virulence genes; P. lacertae is considered a nonpathogenic parasite of lizards, although
recent data may indicate invasive properties of Proteromonas.24
It is very likely that genome sequencing and analysis of common STs such as ST1–ST4 is ongoing while this article is being written and eventually studies should include transcriptomic profiling of
isolates from symptomatic and asymptomatic carriers to investigate gene expression in these STs that
appear to account for more than 90% of all human Blastocystis.
4.4 Diagnosis and Typing
Similar to other intestinal parasites, the life cycle of Blastocystis includes a stage where the organism
is passed from carriers by stool. Traditional parasitological methods such as microscopy of permanently stained fixed fecal smears or fecal concentrates enable the detection of Blastocystis to some
extent46; however, application of techniques such as short-term XIVC and especially nucleic acid–
based methods, such as PCR and sequencing directly applied to fecal DNA template, has made it clear
that Blastocystis carriage is much more common than previously anticipated (unpublished observations).47 This boils down mostly to the fact that most parasitological methods are significantly less
sensitive than PCR.46
Short-term XIVC has been employed in a multitude of cases and surveys4,46,48–54 and uses simple and
inexpensive media such as Robinson’s or Jones’ medium.55,56 The sensitivity of XIVC is superior to
microscopy of fecal concentrates but inferior to real-time PCR.53,54 An advantage of XIVC is that isolates
of interest can be maintained by subculturing for months/years and potentially be cryopreserved in liquid nitrogen or submitted to efforts aiming toward axenization; however, there is currently no standard
protocol available for axenizing Blastocystis cultures. Even xenic cultures are useful for sequencing of,
for instance, complete SSU rRNA genes or mitochondrial genomes,44 whereas only DNA from axenic
cultures is appropriate for complete nuclear genome sequencing; a few axenic strains are available in the
ATCC collection.
The first publication on a diagnostic PCR for Blastocystis came into being in 200657 and was partly
inspired by the emerging trend toward screening fecal DNAs for common intestinal protozoa by PCR as
a supplement to or even as a substitute for microscopy of fecal concentrates. Primer design was based on
sequences available at the NCBI database at that particular time; subsequent analysis of newer data has
led to speculation that this PCR assay may exhibit preferential amplification of one ST over another due
to sequence variation in primer annealing sites, which may impair its use in molecular characterization.
Indeed, the extensive intrageneric diversity of Blastocystis has made the design of a diagnostic genusspecific PCR applicable to fecal DNA template challenging.
Three diagnostic real-time PCR (qPCR) assays have been published. One of them58 was validated only
against ST1, ST3, and ST4. Primers and gene target were not given in the paper, and so little is known
about the sensitivity of this assay. The two other assays are intended to target all STs found in humans
(i.e., ST1–ST9) and are based on the SSU rRNA gene. The first of these two assays to be published was
based on SYBR Green–based detection of Blastocystis-specific DNA and enables subsequent subtyping
(see section 4.4.2) of PCR products, which is a clear advantage.53 Disadvantages include the fact that the
PCR product is quite large (320–342 bp depending on ST) and that the specificity of the assay is only
95%. The second assay is based on TaqMan technology and characterized by 100% specificity.54 The use
of real-time PCR in large-scale surveys will assist in identifying whether the development of symptoms
is related to infection intensity by simple analysis of cycle threshold (Ct) values for individual samples.
XIVC sensitivity ranged between 52% and 79% compared to these real-time PCR assays.53,54 Previously,
XIVC was found to have a sensitivity of 89% compared to a conventional PCR,46 but, importantly, conventional PCR relies on visual evaluation of PCR results, and the conventional assay was based on primers amplifying a relatively large PCR product (~550–585 bp), which was suitable for sequencing and ST
identification, but too large to be relevant for diagnostic PCR, especially in situations where fecal DNAs
are of suboptimal purity. An estimate of the number of copies of rDNA in one Blastocystis cell is not
available, although it may lie somewhere between 10 and 10054; Denoeud et al.38 obtained at least 20
copies by full genome sequencing of ST7.
Recently, Blastocystis was detected and subtyped by screening metagenomic species (MGS) data
generated during a study of the intestinal microbiome of close to 400 Danish and Spanish patients and
healthy individuals by the MetaHIT Consortium (unpublished observations). Of all the genes in the gene
catalog, 2063 blasted to Blastocystis genes. Of these, 1737 were found in MGS clusters. Five clusters had
an overrepresentation of Blastocystis genes. Of these, four could be annotated as ST1, ST2, ST3, and ST4
on the basis of the mitochondrial rps12; it was found that 81/396 (20.5%) study individuals were positive for Blastocystis, and of these, 14 had ST1; 14, ST2; 17, ST3; and 36, ST4. Hence, this is a novel and
potentially very useful way of extracting data on Blastocystis from pools of data already available59 and
has the added benefit of enabling the analysis of Blastocystis in the context of the bacterial microbiota.
4.4.1 In Vitro Isolation of Blastocystis
Clinical Blastocystis isolates are easily established in culture by inoculation of fecal sample in Jones’ or
Robinson’s medium. There are currently no protocols enabling consistent axenization of Blastocystis;
an axenic Blastocystis culture is a culture in which there are no other metabolically active organisms
present than Blastocystis. This means that there are only few axenic reference strains available in the
ATCC collection and elsewhere. Axenic cultures of Blastocystis are essential to a number of research
areas that eventually will lead to knowledge on clinical significance and management of the parasite.
The complete genome is available for Blastocystis sp. ST7, but not for other STs, including STs common
in humans. Part of this is due to these STs not being available as axenic strains and/or that enough DNA
cannot be retrieved from available isolates. This precludes, for instance, biochemical studies, production
of mono- and polyclonal antibodies, and generation of molecular data for studies of comparative genomics and proteomics, including the identification of virulence factors and other effector proteins; attempts
to investigate potential drug targets and therapies are thereby also hampered. Clearly, a standardized and
effective protocol for axenization of Blastocystis is warranted.
4.4.2 Subtyping
Currently, the molecular epidemiology of Blastocystis can be studied at three levels: at ST level, at 18S
allele level, and at sequence type (SQT) level by MLST.
Subtyping, that is, the assignment of ST numbers to Blastocystis identified in a given host, has been
approached by mainly two methods: one method uses seven pairs of diagnostic sequence-tagged site
(STS) primers targeting ST1–ST7 obviating the need for sequencing,60 and the other method relies on
Biology of Foodborne Parasites
sequencing of SSU rDNA amplicons, for instance, by bar coding.61 These two methods were recently
compared.62 While the STS primers appear highly specific even when applied directly to fecal DNA
template, some of the primer pairs are limited by low sensitivity. This is most evident in the case of
ST4. This ST comprises two major clades,44,49 one of which is much more common in humans than the
other. The STS primer pair targeting ST4 may miss the more common clade, hence underestimating the
prevalence of ST4.62 This, however, may not be the case, since one would expect that there would be quite
a few untypable cases of Blastocystis in studies that have employed these primers on DNAs extracted
from cultures, and this appears not to occur. So far, the STS primers have been used in over 20 surveys,
especially in Asian and Middle Eastern countries, where reports of ST4 are surprisingly rare, and so,
rare reports of ST4 coincide with the use of insensitive ST4 primers. The gene targets of the STS primers remain unknown; substantial sequence variation in the region amplified by some of the primers has
been detected,62 although there has been no analysis of genetic diversity at the primer annealing sites
themselves. It should, moreover, be noted that STS primers are not available for ST8 and ST9 or for
any other STs for that matter, which means that screening animal samples by STS primers is of limited
value, unless only specific STs belonging to ST1–ST7 are being looked for. It should be noted, however,
that while the STS primers are characterized by maximum specificity when used on fecal DNAs from
humans, the specificity of the primers when used on fecal DNAs from animals remains unknown.
The second method relies on analysis by sequencing of SSU rDNA PCR products amplified by “panBlastocystis” primers, for example, by bar coding. Bar-code primers (RD5/BhRDr) amplify the 0.6
5′-most kb of the SSU rRNA gene, and phylogenetic analysis has demonstrated that this region is a valid
genetic marker of complete SSU rDNA sequences.61 The drawbacks compared to the STS method are
that sequencing is needed and that mixed ST colonization may be difficult to decipher. On the other
hand, bar coding enables a more subtle analysis, namely, 18S allele analysis. A public database is available at, which includes a sequence depository for bar-code sequences and
sequences obtained by MLST (see Section 4.5.1) along with a BLAST facility where individual or bulk
fasta files can be uploaded for quick allocation of ST number, hence obviating the need for phylogenetic
analysis.44,63 An 18S allele analysis is a valid indicator of intra-ST genetic variation,44 and to date, more
than 35 18S alleles for ST3 have been identified, whereas the number of 18S alleles for ST4 and some
other STs remains much more limited; however, some of the allelic variation is due to cloning, and intrastrain SSU rDNA polymorphism has been reported.38
Other methods used for SSU rDNA-based subtyping include ST-specific single-nucleotide polymorphism by pyrosequencing.64
4.4.3 Impact of Sampling and Sample Processing on
Blastocystis Detection and Subtyping
Methodological issues other than those pertaining to the choice of subtyping method may influence our
ability to accurately identify the distribution of Blastocystis STs in any given population. For instance,
sometimes Blastocystis is subtyped directly from fecal DNA template, sometimes from DNA template
prepared from cultured isolates; there is evidence that XIVC may favor one ST over another in cases of
mixed ST colonization,65 and it is also possible that different strains (e.g., some animal vs. human strains)
may have different requirements in terms of growth conditions, including temperature and nutrients;
however, in some labs, short-term XVIC appears to prevent differential growth of STs affecting the
results obtained.29
Storage conditions before sample processing may be another variable potentially influencing our ability to detect Blastocystis. Samples are often collected in locations far from the lab and may be kept under
varying conditions such as on cold/freeze storage, at ambient temperature, or in preservatives such as
lysis buffer or ethanol.66,67
DNA extraction can be performed in many ways, and differences in quality (purity, presence of
inhibitors) may obviously impact our ability to amplify and sequence Blastocystis targets; here, the
size of the PCR product is also important, since short PCR products may be amplifiable even from low-­
quality DNA template, while longer products (e.g., 600 bp used for bar coding) may be more difficult to
obtain (unpublished observations). Also, the amount of competing DNA template in, for instance, DNA
extracted directly from feces may influence PCR results.
Although clearly advantageous, it may prove much more difficult to standardize sampling and sample
­processing than to standardize subtyping methods. Suffice to say, direct screening of fecal DNA templates by
PCR (preferably real-time PCR or a conventional PCR amplifying a “short” PCR product) with ­subsequent
bar coding of positive samples may be considered “state of the art” due to high diagnostic sensitivity. It has
the added benefit of automatically assigning sequences to an ST number and even 18S alleles when using the
website and should be feasible in most laboratories.
4.5 Epidemiology and Molecular Epidemiology
To give a summary of the prevalence of Blastocystis in various human cohorts and in different countries
is beyond the scope of this chapter, primarily because differences in prevalence figures may equally
well reflect differences in diagnostic modalities as differences in factual prevalence. Suffice to say, in
many countries, especially third world countries, carriers are probably more abundant than noncarriers.
It should be noted though that the prevalence in even highly industrialized countries may amount to
somewhere between 0% and 50% depending on cohort and age (unpublished observations).
In contrast to a parasite such as Dientamoeba fragilis, it appears that Blastocystis prevalence increases
by age or at least that Blastocystis is a lot less common in children than in adults (unpublished observations), suggesting a different mode of transmission and/or a tendency toward chronic colonization.
The conundrum as to whether Blastocystis-positive adults with symptoms are in their first colonization
The fact that Blastocystis is so common gives rise to many interesting questions: Which are the determinants of Blastocystis colonization? Which are the exposures? Are we all repeatedly exposed but only
some susceptible? Is Blastocystis carriage an indicator of a certain behavior (differential exposure), and
can this explain any age-related differences in prevalence? Do humans acquire Blastocystis directly from
other humans (direct contact), from animals, or from the environment? We may already be able to give
a partial answer to the last question, but this requires an understanding of the molecular epidemiology
of Blastocystis.
4.5.1 Molecular Epidemiology of Blastocystis
The host spectrum of Blastocystis appears immense29 and may expand even more as a consequence of
additional sampling. Since there is a considerable overlap of STs between human and nonhuman hosts,
animals might constitute a reservoir for human Blastocystis.
MLST analysis of Blastocystis is currently based on analysis of loci in the MLO genome 44; mitochondrial DNA is especially useful in MLST analysis due to its haploid structure, hence bypassing
the problems related to sequence heterozygosity seen, for instance, in some Giardia MLST loci.68
So far, MLST systems are available for ST3 and ST4, and similar assays for ST1 and ST2 will follow. Analysis of 132 ST3 and ST4 isolates from human and nonhuman primates (NHPs) recently
revealed dramatic differences in intra-ST diversity. No less than 58 SQTs were detected among 81
ST3 samples, while only 5 SQTs were found among 50 ST4 samples belonging to the common genotype.44 ST4 samples obtained from Denmark, England, and Nigeria shared the same SQT. Moreover,
an ST4 strain infecting a guinea pig was found identical to strains infecting humans, and so the ST4
MLST system fails to differentiate between human and rodent strains. It should be kept in mind,
however, that these strains may in fact be identical and that guinea pigs may constitute a reservoir
for human ST4 (or vice versa). Meanwhile, MLST proved a potent tool to distinguish between ST3
strains. While some NHP ST3 strains clustered among human ST3 strains, most NHP ST3 strains
belonged to specific distinct clades. Interestingly, a couple of human ST3 strains were found in the
NHP clades, indicating possible zoonotic transmission, and indeed one of the human strains was
found in a sample from a zookeeper.
Biology of Foodborne Parasites
While MLST, at least for ST3, appears to be a very useful tool for investigating patterns of transmission, the information obtained by the much more simple 18S allele analysis can be an extremely
cost-effective tool. A recent study of Blastocystis in NHPs representing 30 genera showed that NHPs
typically host the same STs as humans, apart from the fact that ST4 is rare in NHPs, while ST5 and ST8
are rare in humans but common in NHPs. However, despite the commonness of ST1 and ST3, it was
seen that many of the ST3 alleles found in NHPs are alleles that are not found in humans, and the same
holds true for ST1.38 A large overlap, however, appears to be present for ST2, but only MLST can confirm
whether or not NHP ST2s are indeed identical to human ST2s.
ST3 appears to be much more common in humans than in other hosts. However, some common synanthropic hosts such as cats and dogs have been studied only to a limited extent. Studies of dogs in
Thailand have revealed reasonably low-prevalence figures,69,70 while another study71 found that 70.8% of
72 dogs and 67.3% of 52 cats studied in Brisbane were positive by simple wet mount analysis, suggesting
a high level of colonization in these animals in this particular region, consistent with recent data coming
out from Queensland.48 In Chile, a large study of diarrhoeic dogs (n = 972) and cats (n = 230) showed
prevalences of 36% and 37%, respectively.72 Dogs appear to be natural hosts of ST1–ST3 and cats of ST1
and ST428,48; however, very few cats and dogs have been surveyed by subtyping, and 18S allele or MLST
analysis of Blastocystis will reveal whether human Blastocystis may stem from such a reservoir.
A comprehensive list of Blastocystis sp. STs found in humans and major groups of nonhuman mammalian and avian hosts is given in Table 4.2. Some host species have been studied to a limited extent
and yet can be shown to be natural hosts of many STs, for instance, lemurs or camels.38 On the contrary,
some host species have been sampled extensively, but the typical range of STs seen appears limited; one
such example is cattle, which have been sampled in many geographic regions: ST10 appears extremely
common along with—to some extent—ST5, while other STs (ST1, ST3, and ST14) appear rare.
Major differences between humans and NHPs can be identified (Table 4.2): ST4 appears extremely
rare in NHPs, while ceboids (new world monkeys) are common hosts of ST8, and ST5 is common in
apes. All STs found in humans, apart from ST9, are also found in NHPs. In contrast, NHPs harbor STs
not yet found in humans: Lemurs appear to host ST10; ST13 was found in cercopithecoids and ST15 in
nonhuman hominoids38; moreover, there is evidence in GenBank (acc. no.: JX159284) of a novel ST from
a gorilla phylogenetically related to reptilian isolates.
In terms of the zoonotic potential of Blastocystis, it may be concluded based on currently available data
that human colonization due to ST1–ST3 (and possibly ST4) may be a consequence primarily of humanto-human transmission, whereas STs found in humans and not belonging to the usual spectrum (ST1–
ST4) may very well represent cases of zoonotic transmission. This conclusion is based mainly on three
observations: (1) STs colonizing humans are mostly different from those colonizing livestock a­ nimals29;
(2) ­analysis of intra-ST diversity (18S allele and MLST analysis) enables us to compare strains belonging
to the same ST but found in different hosts, and, so far, only few examples of potential zoonotic transmission (i.e., shared 18S allele or MLST SQT) have been found44; and (3) some STs may be rare in humans
and mainly found in individuals who have a history of close contact to animals.38,44,61 However, pets such
as cats and dogs appear to a large extent to be colonized by STs seen in humans,48 and 18S allele or MLST
analysis of Blastocystis from such animals is warranted to identify whether such animals constitute a
reservoir for strains colonizing humans.
Studies of intra-ST genetic diversity of Blastocystis have so far enabled us to predict the likelihood of
zoonotic transmission of individual STs. Moreover, the restricted reservoir and the homogeneity of ST4
indicate that this ST has entered the human population only recently compared to other common STs
found in humans. The extensive intragenetic diversity in ST3 on the other hand may bear witness to a
long history of coevolution with humans3,44; this situation may hold one of the keys to understanding the
ST-dependent differences in clinical outcome of colonization by these STs as reported.4,49,73
4.5.2 Linking Molecular Epidemiology to Clinical Presentation
Numerous studies coming out from many different geographic regions have pursued a link between
STs and clinical presentation. On a global scale, no clear trends have been identified, and this may be
explained by one or more of the following reasons: (1) Blastocystis or any ST thereof is not consistently
linked to the development of disease; (2) Blastocystis may cause symptoms initially in previously
naïve patients, while symptoms wear off during chronic colonization possibly due to acquired immunity (in which case the pathogenicity of Blastocystis should be studied longitudinally, i.e., over time);
(3) symptoms caused by Blastocystis may be vague and difficult to define, what may be considered
“normal” by one individual may be considered an ailment in others, which is why the use of standardized, detailed questionnaires are useful in clinical research; (4) different methodologies have been
used for subtyping potentially masking associations; and (5) the variable geographic distribution of
Blastocystis STs may imply variable prevalence of potentially pathogenic strains. The latter situation is
exemplified by the fact that ST4 has been associated with diarrhea in independent studies in Europe,49,73
and in a recent study,4 ST4 was almost statistically associated with irritable bowel syndrome (IBS, see
­section 4.6). ST4 appears virtually absent in most regions outside Europe, and if ST4 is supposedly
linked to disease, while common STs such as ST1–ST3 are not, this may explain why studies coming
out from non-European countries mostly fail to find any consistent association between ST and disease.
The lack of intra-ST genetic variation in ST4 supports the theory that this ST began to colonize the
human population only recently compared to ST1–ST3, which again may explain why ST4 is much less
common in most non-European countries.3,4 This ST has been found in rodents such as guinea pigs and
rats; these animals have been considered a potential reservoir of human infection, and while this may
hold true, many rodents appear not to be carrying Blastocystis,29 and moreover, ST4 has been found in
nonrodent animal hosts.38
One remarkable exception to the rule that no clear association between STs and clinical phenotype is
detected outside Europe is a recent study by Ramirez et al. (submitted for publication) from Colombia,
in which ST1 was found only in patients with non-IBS intestinal symptoms and ST2 was found only in
patients with IBS and ST3 in asymptomatic individuals. The 18S alleles detected for ST1 were identical
to those detected in ST1 isolates in Europe, where ST1 appears not to be linked to disease.
Scanlan and Marchesi47 speculated on a potential age-dependent ST distribution, finding ST4 among
the young and ST3 and ST2 among the elderly; however, any association between age and ST could not
be identified by reviewing data coming out from our own lab (unpublished observations).
4.6 Pathogenesis and Clinical Features
Some facts support the hypothesis that asymptomatic carriage is normal and that Blastocystis may generally not be linked to acute onset of symptoms: the parasite is extremely common and apparently not
linked to outbreaks, and invasive properties have not been convincingly demonstrated, at least not in
humans where colonoscopy is usually normal.48,74
As mentioned, the large degree of diversity in human ST3 points toward a relatively long history of
coevolution,44 and humans may have adapted to this ST, while other STs may be “new” to the human
host and it is tempting to speculate that they, therefore, may be influencing intestinal homeostasis or even
contribute directly to the development of symptoms. One such example could be ST8, which is usually
a parasite of NHPs and only rarely found in humans. This ST was the only potential pathogen identified in stools from a Danish woman hospitalized for 3 days due to diarrhea and fever after returning
from vacation in Indonesia.52 Two courses of metronidazole were not able to eradicate the parasite, and
successful eradication was verified by PCR analysis of multiple fecal samples after a 10-day course of
trimethoprim + sulfamethoxazole (co-trimoxazole) treatment, after which diarrhea, bloating, flatulence,
and abdominal pain disappeared.
Although the scientific literature abounds with reports on symptoms potentially related to Blastocystis,
these symptoms are nonpathognomonic and potentially attributable to other intestinal pathogens
transmitted fecally–orally as well as to other conditions such as IBS. For instance, a parasite such as
D. ­fragilis often colonizes individuals who are also positive for Blastocystis,52,75,76 but generally, analysis
for D. ­fragilis is rarely included in a microbiological work-up.
A parameter such as infection intensity has been thought to play a role in the development of symptoms. And while a relationship between infection intensity and symptoms may appear plausible, the
methods by which this has been investigated should be questioned. For instance, it has been customary
Biology of Foodborne Parasites
to define Blastocystis colonization as having more than five organisms per visual field by microscopy.
Such an approach may have very limited value in the light of day-to-day variation in parasite shedding5
and variable sensitivity of microscopy-based methods. The real-time PCR assays recently developed53,54
will standardize Blastocystis detection as well as allow exploration of any role for infection intensity in
A couple of case reports warrant some attention: in 2010, Vogelberg et al. published a report on a
case where Blastocystis sp. ST2 was apparently associated with urticaria and flatulence in a 20-yearold male returning to Germany from China.77 Full blood count including eosinophil count, erythrocyte
sedimentation rate, C-reactive protein, cryoglobulins, circulating immune complexes, C3, C4, CH 100,
C1-INH, IgE, ANA, MAK, TAK, TRAK, immunoglobulins (IgA, IgM, and IgG), and serological investigations (EBV, Coxsackie B1–B6, A9, ECHO virus pool, TSH, T3, and T4) were all within the normal
range. Radio allergen sorbent testing revealed a sensitization against house-dust mite and tree and grass
pollen. Hence, malabsorption, endocrinological, rheumatological, and autoimmune diseases could be
ruled out. A microbiological and parasitological examination of three consecutive stools revealed no
bacterial enteropathogens, no enteroviruses, and no intestinal protozoa (including microsporidia and
D. fragilis) apart from Blastocystis, which was present in all three samples; in particular, no bacterial
toxins could be detected. Multiple courses of different drug regimens were completed, after each of
which clinical and microbiological evaluations were performed. Colonoscopy revealed heavy colonization by vacuolar stages of Blastocystis and discrete unspecific mucosal inflammation, while histological
analysis produced no evidence of invasion. Blastocystis was eventually eradicated by a combination of
metronidazole and paromomycin with concurrent resolution of both intestinal and urticarial symptoms;
monotherapy with metronidazole, sulfamethoxazole and trimethoprim, and paromomycin had proven
ineffective for eradication.
Other reports linking Blastocystis to urticaria include studies by Pasqui et al.78 and Zuel-Fakkar et al.19
Importantly, Blastocystis-related urticaria is not associated to ST2 only.19 This may indicate that this
symptom may be dependent on factors other than ST. Virulence may, for instance, be “switched on” by
regulation of gene expression for reasons that are not yet understood. It may also be that host factors are
determinants of clinical outcome. It has been speculated that Blastocystis-mediated urticaria may be due
to parasite antigen activation of specific clones of Th-2 lymphocytes to release cytokines (IL3, IL4, IL5,
IL13) with consequent IgE production and mast cell activation.78
Given the prevalence of Blastocystis, colonization of patients with colonic/rectal ulcers may be a quite
common finding. Sometimes, Blastocystis has been associated with pathological lesions in the distal
gut79,80; however, since usually no or a very few details are given regarding potential differential diagnostic considerations, such as the possibility of entamoebiasis or bacterial colitis, linking Blastocystis
to invasion may be misleading, since the parasite could potentially end up in tissue lesions generated by
coinfecting invasive pathogens, hence being a secondary finding.
With regard to colitis, a few studies have surveyed the prevalence of Blastocystis in patients with
inflammatory bowel disease (IBD). Cekin et al.81 found that Blastocystis was significantly more common in patients with ulcerative colitis (UC) than in a control group, while patients with Crohn’s disease (CD) and IBS were not significantly more frequent carriers than controls. Recent studies by our
group, however, point toward a completely different scenario: in a study of 100 Danish IBD patients and
96 controls, Blastocystis was found significantly more often in controls (19%) than in patients with IBD
(5%) (p < 0.05), while D. fragilis was found in more or less equal numbers, 15% and 13%, respectively.
Blastocystis and D. fragilis were more predominant in patients with UC in remission (19% and 33%)
compared to patients with active disease (0% and 5%, p < 0.05). Remarkably, Blastocystis was found in
only 1/42 (2%) of patients with CD. Among healthy controls, Blastocystis and D. fragilis colonization
was not associated with an elevated level of fecal calprotectin, a surrogate marker of intestinal inflammation.82 Adding support to these trends, Blastocystis-specific DNA could not be amplified from fecal
specimens from 20 Spanish patients with CD, while 17% of 127 patients with UC and 24% of 72 healthy
individuals were found to be colonized using metagenomic analysis (unpublished data).
Analysis of metagenomic data using fecal DNA furthermore led to the observation that Blastocystis
is rare in individuals in whom Bacteroides is the predominating intestinal microbiota component, while
common in individuals with microbiota predominated by Ruminococcus–Prevotella (unpublished
observations). The implications of this so far are unclear, but again support the incentive for investigating the role and activity of Blastocystis in a microbiological and ecological context. Interestingly, a
pyrosequencing study of fecal samples from 40 twin pairs showed that gastrointestinal microbial profiles
vary with IBD phenotypes and that the microbial compositions of individuals with CD differed from
those of healthy individuals but were similar between healthy individuals and individuals with UC83; Qin
et al.,84 however, were able to separate not only UC patients from CD patients but also healthy individuals from IBD patients, based on microbiota profiling. Butyrate and other SCFAs, which are produced
by groups of intestinal bacteria (especially members of the phylum Firmicutes such as Lachnospiraceae
and Ruminococcaceae) and have a number of functions essential to colonic health and immune function, not only are effective antimicrobials but also regulate cells of both the innate and adaptive immune
systems.85 SCFAs have the ability to inhibit the proliferation and activation of T cells and prevent adhesion of antigen-presenting cells, and butyrate in particular exerts potent anti-inflammatory activity by
inhibiting NF-κB signaling.85 Significant reduction in the abundance of bacteria involved in butyrate and
propionate metabolism, including Ruminococcus bromii et rel., Eubacterium rectale et rel., Roseburia
sp., and Akkermansia sp., is among the markers of dysbiosis in UC.84 Joossens et al. showed a lower
prevalence of butyrate-producing bacteria (e.g., Faecalibacterium prausnitzii) in patients with CD compared to their unaffected relatives and healthy controls.87 Vermeiren et al.88 demonstrated depletion in
butyrate-producing microbial communities in samples from UC patients compared to healthy volunteers
using an in vitro dynamic gut model. These are only a few of the many observations and considerations
that highlight the necessity and relevance of characterizing the microbiota structure, diversity, and function that accompany Blastocystis colonization along with measurements of butyric acid, calprotectin,
and other biomarkers of intestinal homeostasis and inflammation in order to identify growth requirements and potential interaction with the host and surrounding microbiota.
4.6.1 Blastocystis and Intestinal Eukaryotic Microbiota
Apart from targeted detection and characterization of the parasite, molecular data on Blastocystis have
been obtained in studies aiming to characterize the overall structure of the fecal eukaryotic microbiota.
Scanlan and Marchesi47 used culture-independent phylogenetic interrogation of cloned and sequenced 18S
and ITS PCR products amplified from fecal DNA template from 17 healthy individuals and analyzed by
denaturing gradient gel electrophoresis (DGGE) to show that the eukaryotic diversity of the human gut
is low, largely temporally stable, and predominated by different STs of Blastocystis. A total of 14/17 individuals were shown to be colonized and harbored STs belonging to ST2, ST3, and ST4, and importantly,
no cases of mixed STs were detected by this—in theory extremely sensitive—method.
A somewhat similar approach was taken by Hamad et al.89 who reported on the eukaryotic diversity in
a single fecal sample of a Senegalese male, in whom Blastocystis was found. Pandey et al.34 used cloning
of SSU rDNA PCR products amplified from template extracted from fecal samples from infants and their
respective mothers and found that infants were not colonized by microeukaryotes, while their mothers
were typically colonized by Blastocystis sp., Saccharomyces sp., and Candida albicans.
Surprisingly, these are the only examples of studies where the total eukaryotic component of the
human intestinal microbiome has been addressed to date, despite the fact that some microeukaryotes
such as Blastocystis, Dientamoeba, and yeasts are common constituents of the commensal flora and
despite the current hyperbole surrounding the human intestinal microbiome.90
While popular working hypotheses foresee a link between ST and virulence or infection intensity
and severity of symptoms, it may very well be that other factors may be responsible for pathogenicity
attributable directly or indirectly to Blastocystis. Scanlan91 hypothesized that Blastocystis may select for
bacterial virulence-related traits that enable grazing resistance such as biofilm formation and survival
within the host when ingested. While predation may be a feature of some protozoa, phagocytosis by
Blastocystis has not yet been convincingly demonstrated. However, it is also possible that Blastocystis
influences the surrounding microbiota in other ways, for example, by secreted proteases,92 and thereby
may be indirectly selecting for certain bacterial phenotypes.
Of tremendous interest is also the question of the determinants of Blastocystis colonization. For
instance, it has been speculated that Blastocystis might be a predictor of a certain type of microbiota,59
Biology of Foodborne Parasites
which may mean that Blastocystis is dependent on certain bacterial communities for colonization,
or might select for such particular communities. Based on analysis of the intestinal microbiome,
humans can be more or less successfully stratified into enterotypes based on relative abundance of
mainly Bacteroides, Prevotella, and Ruminococcus.93 As already mentioned, recent data suggest
that Blastocystis colonization appears rare in individuals dominated by Bacteroides, and interestingly, the intestinal microbiota of IBS patients appears to be predominated by bacteria other than
4.6.2 Blastocystis and Irritable Bowel Syndrome
IBS affects 16% of the Danish adult population (Krogsgaard, personal communication). Given the fact
that the disease is so common and with no obvious etiology, it is hardly surprising that common parasites
such as Blastocystis and D. fragilis have been subject to scrutiny in these patients as potentially contributing factors. Association between Blastocystis carriage and IBS has been proven in some studies75 but
clearly needs further investigation. Poirier et al.92 recently gave an account of the putative pathogenic
mechanisms by which Blastocystis might contribute to IBS. A recent study from the United Kingdom
analyzed the distribution of STs in fecal samples from patients referred to IBS clinics and found a preponderance of ST4 compared to samples submitted from patients with no explicit history of IBS.4
Individuals diagnosed with IBS are often reported to have a much higher infection rate with Blastocystis
than controls—often twice as high or more.95–98 Moreover, many of the symptoms linked to Blastocystis
infection are very similar to those used to diagnose some types of IBS (diarrhea, vomiting, abdominal
cramps, and bloating). This suggests either that Blastocystis colonization should be a differential diagnosis or that in some cases of IBS, Blastocystis may be the causative agent. As already indicated, it could
also simply mean that the microbiota of the IBS gut means that Blastocystis colonizes more efficiently.
As an intestinal parasite, infection with Blastocystis means exposure to fecal organisms, and so it is perhaps more accurate to say that the data only support a link between fecal exposure and IBS.
A diagnosis of IBS should only follow exclusion of other potential causes of the symptoms, functional
as well as microbiological. IBS is a syndrome with several different forms and diagnosis currently uses
the Rome III criteria: “Recurrent abdominal pain or discomfort at least 3 days per month in the last
3 months associated with 2 or more of the following: (1) improvement with defecation, (2) onset associated with a change in frequency of stool, and (3) onset associated with a change in form (appearance)
of stool.” The “change in frequency” can mean diarrhea (IBS-D) or constipation (IBS-C) or a mixture
of the two (IBS-M), yet in studies of Blastocystis in IBS patients, these variants are not differentiated.
The problem of diagnosis is highlighted by the study of Giacometti et al. from Italy.97 They found
a significant difference (p ≤ 0.01) between the prevalence of Blastocystis in IBS patients (15/81) and
those with other gastrointestinal complaints (23/307). However, at a 6-month follow-up, 53/72 returning
patients no longer met IBS criteria, which would not be expected for a chronic disease.
The interpretation of Blastocystis/IBS associations is complicated by the existence of nine STs in
humans, and only three studies have investigated a link between Blastocystis STs and IBS, with different
results. In Pakistan96 and Egypt,99 the studies found ST1 to be significantly more common in IBS patients
than in controls. In contrast, in the United Kingdom, ST4 was more common and ST1 less common in
patients from IBS clinics, but the differences did not reach statistical significance. There was a difference
in methodology (STS vs. sequencing) among other variables between the studies. So, the potential link
between IBS and Blastocystis STs needs further investigation, and it may be particularly important to
identify the variant of IBS in investigating this relationship.
4.6.3 Blastocystis Animal Models
For many diseases, animal models were essential in establishing the role of the pathogen and in principle
should be a way of investigating the role of Blastocystis in disease. However, to date such studies have not
proven particularly helpful. For example, experimental infections of laboratory rodents have led to observations that are not made in colonized humans, such as tissue invasion100 and increased oxidative stress.101
In vitro studies using axenic Blastocystis have demonstrated induction of cytokines, contact-mediated
apoptosis, and barrier disruption, but there is no evidence that these occur in vivo. Appropriate controls
are also needed—for example, when infecting experimental animals with Blastocystis from cultures
growing with bacteria, animals must also be infected with the bacterial flora alone to be certain that
Blastocystis is responsible for any effects observed.102
4.6.4 In Silico Prediction of Blastocystis Pathogenicity by Genome Analysis
Effector proteins or virulence factors of infectious pathogens are likely involved in cell adhesion, colonization, invasion, and host immune evasion and may include surface proteins as well as secreted proteins,
such as proteases, signaling molecules, and molecules involved in quorum sensing.103 Analysis of the
nuclear genome of Blastocystis ST7 has led to some interesting observations.38,92 The putative secretome of Blastocystis includes fucosidase, hexosaminidase, and polygalacturonase, which may be able
to degrade host glycoproteins. Genes encoding major classes of proteolytic enzymes have been identified in the genome, including serine, aspartic, and cysteine proteases and metalloproteases. Of the 66
proteases identified, 18 were predicted to be secreted, and genes involved in protease maturation have
also been identified (e.g., asparaginyl endopeptidase [legumain]). Several protease inhibitors have been
identified, some of which may inhibit host proteases or defend Blastocystis from nonspecific digestive
enzymes. Blastocystis might be capable of secreting toxins or macrolides, for instance, by a putative type
1 polyketide synthase gene potentially acquired by HGT.
It should be noted that these are in silico observations. Some species of, for instance, Entamoeba are
in possession of similar virulence genes, but it appears that they are not expressed; the mere presence of
virulence genes does not necessarily imply that these genes are expressed or active, and hence the existence of potential virulence markers should not necessarily equate with pathogenicity per se.
4.7 Treatment and Prevention
One of the major limitations in Blastocystis research is the absence of convincing randomized placebocontrolled treatment (RCT) studies that could assist in exploring whether any clinical improvement is
related to microbiological cure. The absence of such studies may stem from our current inability to
eradicate the parasite. Few RCT studies have been carried out,104–106 and studies undertaken so far can
be criticized mainly for (1) not reporting on diagnostic method(s) used, (2) using insensitive diagnostic
methods for postintervention microbiological follow-up, (3) application of diagnostic criteria of limited
relevance, (4) lack of testing for D. fragilis, (5) lack of implementation of good manufacturing practice,
(6) lack of consideration on sample sizes, (7) lack of definition of inclusion criteria, and (8) failure to
conduct thorough longitudinal follow-up on clinical and microbiological status.
Many of the drug regimens used for treating Blastocystis and reported in the scientific literature were
recently summarized107; of particular interest is that a widely applied agent such as metronidazole may—
when used in monotherapy—have limited or no effect in terms of eradicating Blastocystis. Although
clearly efficient in vitro,52 limited in vivo efficacy may not be surprising given the fact that Blastocystis
is a parasite of the colon, while metronidazole is more or less completely absorbed in the small intestine.
The combination of metronidazole and paromomycin has proven efficient in a few cases,77 although it is
clear that treatment failure even by this combination occurs108 and it also remains unclear whether paromomycin alone would be sufficient.
Again, it should be emphasized that Blastocystis is related neither to fungi nor to protozoa, and so it
may prove necessary to identify or develop agents that are targeted specifically at this parasitic stramenopile, for example, by blocking Blastocystis-specific metabolic or reproductive pathways; it would be a
great advantage if the parasite can be selectively targeted.
At present, however, it remains unclear to what extent prevention of Blastocystis colonization should
be prioritized. Some aspects of transmission are still unclear; first and foremost, we do not have a
clear picture of exposure. A few have investigated the presence of Blastocystis in the environment.
Viable cysts have been detected in sewage samples in Scotland, Malaysia, and the Philippines,17,109
and Blastocystis has been detected in samples of river water.110–113 One study has found Blastocystis in
Biology of Foodborne Parasites
leafy vegetables in Saudi Arabia.114 Clearly, more studies are needed on Blastocystis in environmental
samples (rivers, surface water, soil), but also especially in nonprocessed food produce, such as (organic)
vegetables, to identify probably modes and vehicles of transmission and potential points of intervention
where relevant.
4.8 Concluding Remarks and Future Directions
Many factors influence our ability to identify the role of Blastocystis in health and disease. Novel technologies and methodologies are rapidly improving our understanding of the interaction between host and
the intestinal microbiome including the intestinal microbiota’s role in the development of a disease.93,115,116
However, our understanding of the eukaryotic component of the intestinal microbiome is still lagging
woefully behind.90 Studies based on “omics” technologies should quickly advance our knowledge of the
structure and function also of microeukaryotic communities (including Blastocystis) and their potential
ecological interaction with host and surrounding bacterial community.
Comparative genomics and transcriptomic analyses will identify differences in potential virulence
genes and gene expression among STs or subsets thereof. Single-cell genomics117,118 may circumvent
problems related to obtaining sufficient high-quality template from axenic cultures.
Longitudinal epidemiological studies should be undertaken to monitor the incidence of Blastocystis in
different cohorts and whether the initial period of colonization is linked to the development of symptoms
and/or whether symptoms are linked to colonization intensity.
Efficient ways to eradicate Blastocystis should be identified in order to monitor the effect of eradication in symptomatic individuals by randomized controlled treatment studies. In case virulence can be
attributed to Blastocystis, essential proteins should be identified with a view to identifying and/or designing chemotherapeutical agents for specialized and targeted intervention; this may require a combination
of functional and structural genomics to sample the essential Blastocystis structome in order to enable
high-resolution structures of proteins essential to the parasite. Conversely, in case Blastocystis may be
shown to promote a healthy gut environment and/or contribute to a balanced host immune response, the
identification and synthesis of relevant components for use as pre- or probiotics may appear useful.
Deducing the clinical significance of Blastocystis moreover requires further elucidation of aspects
of its life cycle and epidemiology. Although rapidly expanding, our knowledge of genetic diversity and
host specificity is still very limited and should be further elucidated to assist in clarifying transmission
patterns and exposures and to advance investigations into Blastocystis evolution and improve taxonomic
While efforts continue to identify its role in health and disease, we should also try to investigate the
potential of Blastocystis as a proxy or biomarker of certain microbiology phenotypes; for instance, we
should study the potential association between Blastocystis and enterotypes or microbiota function.59
The impact of Blastocystis on host and immunity should be studied in the light of the finding that
Blastocystis is virtually absent in patients with IBDs, at least in some studies, and also in light of the
immunomodulatory functions of other parasites.119
I thank Dr. Graham Clark for commenting on this chapter.
1. Brumpt, E. Blastocystis hominis n. sp. et formes voisines. Bull Soc Pathol Exot 5, 725–730 (1912).
2. Alexeieff, A. Sur la nature des formations dites “Kystes de Trichomonas intestinalis”. C R Soc Biol 71,
296–298 (1911).
3. Clark, C.G., van der Giezen, M., Alfellani, M., and Stensvold, C.R. Recent developments in Blastocystis
research. Adv Parasitol 82, 1–32 (2013). PubMed PMID: 23548084.
4. Alfellani, M.A. et al. Variable geographic distribution of Blastocystis subtypes and its potential implications. Acta Trop 126, 11–18 (2013).
5. Vennila, G.D. et al. Irregular shedding of Blastocystis hominis. Parasitol Res 85, 162–164 (1999).
6. Stenzel, D.J. and Boreham, P.F. Blastocystis hominis revisited. Clin Microbiol Rev 9, 563–584 (1996).
7. Stensvold, C.R., Nielsen, H.V., Mølbak, K., and Smith, H.V. Pursuing the clinical significance of
Blastocystis—Diagnostic limitations. Trends Parasitol 25, 23–29 (2009).
8. Tan, K.S. New insights on classification, identification, and clinical relevance of Blastocystis spp. Clin
Microbiol Rev 21, 639–665 (2008).
9. Stenzel, D.J., Boreham, P.F., and McDougall, R. Ultrastructure of Blastocystis hominis in human stool
samples. Int J Parasitol 21, 807–812 (1991).
10. Vdovenko, A.A. Blastocystis hominis: Origin and significance of vacuolar and granular forms. Parasitol
Res 86, 8–10 (2000).
11. Suresh, K. and Smith, H. Comparison of methods for detecting Blastocystis hominis. Eur J Clin Microbiol
Infect Dis 23, 509–511 (2004).
12. Rene, B.A., Stensvold, C.R., Badsberg, J.H., and Nielsen, H.V. Subtype analysis of Blastocystis isolates
from Blastocystis cyst excreting patients. Am J Trop Med Hyg 80, 588–592 (2009).
13. Stenzel, D.J. and Boreham, P.F. A cyst-like stage of Blastocystis hominis. Int J Parasitol 21, 613–615
14. Zaman, V. The diagnosis of Blastocystis hominis cysts in human faeces. J Infect 33, 15–16 (1996).
15. Zaman, V., Howe, J., and Ng, M. A comparative morphology of Blastocystis hominis cysts with and
without the “fibrillar layer”. Southeast Asian J Trop Med Public Health 26, 801–802 (1995).
16. Moe, K.T. et al. Development of Blastocystis hominis cysts into vacuolar forms in vitro. Parasitol Res 85,
103–108 (1999).
17. Suresh, K., Smith, H.V., and Tan, T.C. Viable Blastocystis cysts in Scottish and Malaysian sewage samples. Appl Environ Microbiol 71, 5619–5620 (2005).
18. Zhang, X. et al. Ultrastructural insights into morphology and reproductive mode of Blastocystis hominis.
Parasitol Res 110, 1165–1172 (2012).
19. Zuel-Fakkar, N.M., Abdel Hameed, D.M., and Hassanin, O.M. Study of Blastocystis hominis isolates in
urticaria: A case-control study. Clin Exp Dermatol 36, 908–910 (2011).
20. Silberman, J.D., Sogin, M.L., Leipe, D.D., and Clark, C.G. Human parasite finds taxonomic home.
Nature 380, 398 (1996).
21. Arisue, N. et al. Phylogenetic position of Blastocystis hominis and of stramenopiles inferred from multiple molecular sequence data. J Eukaryot Microbiol 49, 42–53 (2002).
22. Hoevers, J.D. and Snowden, K.F. Analysis of the ITS region and partial ssu and lsu rRNA genes of
Blastocystis and Proteromonas lacertae. Parasitology 131, 187–196 (2005).
23. Pérez-Brocal, V. and Clark, C.G. Analysis of two genomes from the mitochondrion-like organelle of the
intestinal parasite Blastocystis: Complete sequences, gene content, and genome organization. Mol Biol
Evol 25, 2475–2482 (2008).
24. Maia, J.P., Gómez-Díaz, E., and Harris, D.J. Apicomplexa primers amplify Proteromonas (Stramenopiles,
Slopalinida, Proteromonadidae) in tissue and blood samples from lizards. Acta Parasitol 57, 337–341 (2012).
25. Clark, C.G. Extensive genetic diversity in Blastocystis hominis. Mol Biochem Parasitol 87, 79–83 (1997).
26. Böhm-Gloning, B., Knobloch, J., and Walderich, B. Five subgroups of Blastocystis hominis from symptomatic and asymptomatic patients revealed by restriction site analysis of PCR-amplified 16S-like rDNA.
Trop Med Int Health 2, 771–778 (1997).
27. Stensvold, C.R. et al. Terminology for Blastocystis subtypes—A consensus. Trends Parasitol 23, 93–96
28. Stensvold, C.R. et al. Subtype distribution of Blastocystis isolates from synanthropic and zoo animals and
identification of a new subtype. Int J Parasitol 39, 473–479 (2009).
29. Alfellani, M.A., Taner-Mulla, D., Jacob, A.S., Imeede, C.A., Yoshikawa, H., Stensvold, C.R., and Clark,
C.G. Genetic diversity of blastocystis in livestock and zoo animals. Protist 164(4), 497–509 (2013).
30. Santín, M., Gómez-Muñoz, M.T., Solano-Aguilar, G., and Fayer, R. Development of a new PCR protocol
to detect and subtype Blastocystis spp. from humans and animals. Parasitol Res 109, 205–212 (2011).
31. Fayer, R., Santin, M., and Macarisin, D. Detection of concurrent infection of dairy cattle with Blastocystis,
Cryptosporidium, Giardia, and Enterocytozoon by molecular and microscopic methods. Parasitol Res
111, 1349–1355 (2012).
Biology of Foodborne Parasites
32. Parkar, U. et al. Molecular characterization of Blastocystis isolates from zoo animals and their animalkeepers. Vet Parasitol 169, 8–17 (2010).
33. Petrášová, J. et al. Diversity and host specificity of Blastocystis in syntopic primates on Rubondo Island,
Tanzania. Int J Parasitol 41, 1113–1120 (2011).
34. Pandey, P.K. et al. Molecular typing of fecal eukaryotic microbiota of human infants and their respective
mothers. J Biosci 37, 221–226 (2012).
35. Dunn, L.A., Boreham, P.F., and Stenzel, D.J. Ultrastructural variation of Blastocystis hominis stocks in
culture. Int J Parasitol 19, 43–56 (1989).
36. Stenzel, D.J., Dunn, L.A., and Boreham, P.F. Endocytosis in cultures of Blastocystis hominis. Int
J Parasitol 19, 787–791 (1989).
37. Denoeud, F. et al. Genome sequence of the stramenopile Blastocystis, a human anaerobic parasite.
Genome Biol 12, R29 (2011).
38. Alfellani, M.A., Jacob, A.S., Perea, N.O., Krecek, R.C., Taner-Mulla, D., Verweij, J.J., Levecke, B.,
Tannich, E., Clark, C.G., and Stensvold, C.R. Diversity and distribution of Blastocystis sp subtypes in
non-human primates. Parasitology 140(8), 966–971 (2013).
39. Kittelmann, S. et al. Simultaneous amplicon sequencing to explore co-occurrence patterns of bacterial,
archaeal and eukaryotic microorganisms in rumen microbial communities. PLoS ONE 8, e47879 (2013).
40. Zierdt, C.H. Cytochrome-free mitochondria of an anaerobic protozoan—Blastocystis hominis. J Protozool
33, 67–69 (1986).
41. Stechmann, A. et al. Organelles in Blastocystis that blur the distinction between mitochondria and hydrogenosomes. Curr Biol 18, 580–585 (2008).
42. Tsaousis, A.D. et al. Evolution of Fe/S cluster biogenesis in the anaerobic parasite Blastocystis. Proc Natl
Acad Sci USA 109, 10426–10431 (2012).
43. Wawrzyniak, I. et al. Complete circular DNA in the mitochondria-like organelles of Blastocystis hominis.
Int J Parasitol 38, 1377–1382 (2008).
44. Stensvold, C.R., Alfellani, M., and Clark, C.G. Levels of genetic diversity vary dramatically between
Blastocystis subtypes. Infect Genet Evol 12, 263–273 (2012).
45. Tsaousis, A.D., Stechmann, A., Hamblin, K.A., van der Giezen, M., Pérez-Brocal, V., and Clark, C.G.
2010. The Blastocystis mitochondrion-like organelles. In Anaerobic Parasitic Protozoa: Genomics
and Molecular Biology (C.G. Clark, P.J. Johnson, and R.D. Adam, eds.), Caister Academic Press Ltd,
Norwich, Norfolk, U.K., pp. 205–219.
46. Stensvold, C.R., Arendrup, M.C., Jespersgaard, C., Mølbak, K., and Nielsen, H.V. Detecting Blastocystis
using parasitologic and DNA-based methods: A comparative study. Diagn Microbiol Infect Dis 59, 303–
307 (2007).
47. Scanlan, P.D. and Marchesi, J.R. Micro-eukaryotic diversity of the human distal gut microbiota: Qualitative
assessment using culture-dependent and -independent analysis of faeces. ISME J 2, 1183–1193 (2008).
48. Nagel, R. et al. Blastocystis subtypes in symptomatic and asymptomatic family members and pets and
response to therapy. Intern Med J 42, 1187–1195 (2012).
49. Stensvold, C.R., Christiansen, D.B., Olsen, K.E., and Nielsen, H.V. Blastocystis sp. subtype 4 is common in
Danish Blastocystis-positive patients presenting with acute diarrhea. Am J Trop Med Hyg 84, 883–885 (2011).
50. Stensvold, C.R. and Nielsen, H.V. Comparison of microscopy and PCR for the detection of intestinal
parasites in Danish patients supports incentive for molecular screening platforms. J Clin Microbiol 50,
540–541 (2011).
51. Stensvold, C.R., Arendrup, M.C., Mølbak, K., and Nielsen, H.V. The prevalence of Dientamoeba fragilis
in patients with suspected enteroparasitic disease in a metropolitan area in Denmark. Clin Microbiol
Infect 13, 839–842 (2007).
52. Stensvold, C.R., Arendrup, M.C., Nielsen, H.V., Bada, A., and Thorsen, S. Symptomatic infection with
Blastocystis sp. subtype 8 successfully treated with trimethoprim-sulfamethoxazole. Ann Trop Med
Parasitol 102, 271–274 (2008).
53. Poirier, P. et al. Development and evaluation of a real-time PCR assay for detection and quantification of
Blastocystis parasites in human stool samples: Prospective study of patients with hematological malignancies. J Clin Microbiol 49, 975–983 (2011).
54. Stensvold, C.R., Ahmed, U.N., Andersen, L.O., and Nielsen, H.V. Development and evaluation of a
genus-specific, probe-based, internal process controlled real-time PCR assay for sensitive and specific
detection of Blastocystis. J Clin Microbiol 50, 1847–1851 (2012).
55. Clark, C.G. and Diamond, L.S. Methods for cultivation of luminal parasitic protists of clinical importance. Clin Microbiol Rev 15, 329–341 (2002).
56. Leelayoova, S. et al. In-vitro cultivation: A sensitive method for detecting Blastocystis hominis. Ann Trop
Med Parasitol 96, 803–807 (2002).
57. Stensvold, R., Brillowska-Dabrowska, A., Nielsen, H.V., and Arendrup, M.C. Detection of Blastocystis hominis in unpreserved stool specimens by using polymerase chain reaction. J Parasitol 92, 1081–1087 (2006).
58. Jones, M.S. et al. Detection of Blastocystis from stool samples using real-time PCR. Parasitol Res 103,
551–557 (2008).
59. Stensvold, C.R. Thinking Blastocystis out of the box. Trends Parasitol 28, 305 (2012).
60. Yoshikawa, H. et al. Polymerase chain reaction-based genotype classification among human Blastocystis
hominis populations isolated from different countries. Parasitol Res 92, 22–29 (2004).
61. Scicluna, S.M., Tawari, B., and Clark, C.G. DNA barcoding of Blastocystis. Protist 157, 77–85 (2006).
62. Stensvold, C.R. Comparison of sequencing (barcode region) and sequence-tagged-site PCR for
Blastocystis subtyping. J Clin Microbiol 51, 190–194 (2013).
63. Jolley, K.A. and Maiden, M.C. BIGSdb: Scalable analysis of bacterial genome variation at the population
level. BMC Bioinform 11, 595 (2010).
64. Stensvold, C.R. et al. Blastocystis: Subtyping isolates using pyrosequencing technology. Exp Parasitol
116, 111–119 (2007).
65. Parkar, U. et al. Direct characterization of Blastocystis from faeces by PCR and evidence of zoonotic
potential. Parasitology 134, 359–367 (2007).
66. Malheiros, A.F., Stensvold, C.R., Clark, C.G., Braga, G.B., and Shaw, J.J. Short report: Molecular characterization of Blastocystis obtained from members of the indigenous Tapirapé ethnic group from the
Brazilian Amazon region, Brazil. Am J Trop Med Hyg 85, 1050–1053 (2011).
67. Forsell, J., Granlund, M., Stensvold, C.R., Clark, G.C., and Evengård, B. Subtype analysis of Blastocystis
isolates in Swedish patients. Eur J Clin Microbiol Infect Dis 31, 1689–1696 (2012).
68. Ankarklev, J., Svärd, S.G., and Lebbad, M. Allelic sequence heterozygosity in single Giardia parasites.
BMC Microbiol 12, 65 (2012).
69. Inpankaew, T., Traub, R., Thompson, R.C., and Sukthana, Y. Canine parasitic zoonoses in Bangkok temples. Southeast Asian J Trop Med Public Health 38, 247–255 (2007).
70. Leelayoova, S., Siripattanapipong, S., Naaglor, T., Taamasri, P., and Mungthin, M. Prevalence of intestinal parasitic infections in military personnel and military dogs, Thailand. J Med Assoc Thai 92(Suppl. 1),
S53–S59 (2009).
71. Duda, A., Stenzel, D.J., and Boreham, P.F. Detection of Blastocystis sp. in domestic dogs and cats. Vet
Parasitol 76, 9–17 (1998).
72. López, J., Abarca, K., Paredes, P., and Inzunza, E. Intestinal parasites in dogs and cats with gastrointestinal symptoms in Santiago, Chile. Rev Med Chil 134, 193–200 (2006).
73. Domínguez-Márquez, M.V., Guna, R., Muñoz, C., Gómez-Muñoz, M.T., and Borrás, R. High prevalence
of subtype 4 among isolates of Blastocystis hominis from symptomatic patients of a health district of
Valencia (Spain). Parasitol Res 105, 949–955 (2009).
74. Zuckerman, M.J., Watts, M.T., Ho, H., and Meriano, F.V. Blastocystis hominis infection and intestinal
injury. Am J Med Sci 308, 96–101 (1994).
75. Stensvold, C.R. et al. Blastocystis: Unravelling potential risk factors and clinical significance of a common but neglected parasite. Epidemiol Infect 137, 1655–1663 (2009).
76. Engsbro, A.L., Stensvold, C.R., Nielsen, H.V., and Bytzer, P. Treatment of Dientamoeba fragilis in
patients with irritable bowel syndrome. Am J Trop Med Hyg 87, 1046–1052 (2012).
77. Vogelberg, C. et al. Blastocystis sp. subtype 2 detection during recurrence of gastrointestinal and urticarial symptoms. Parasitol Int 59, 469–471 (2010).
78. Pasqui, A.L. et al. Chronic urticaria and Blastocystis hominis infection: A case report. Eur Rev Med
Pharmacol Sci 8, 117–120 (2004).
79. Janarthanan, S., Khoury, N., and Antaki, F. An unusual case of invasive Blastocystis hominis infection.
Endoscopy 43(Suppl. 2 UCTN), E185–E186 (2011).
80. Lucía, J.F., Aguilar, C., and Betran, A. Blastocystis hominis colitis in a haemophilic patient as a cause of
lower gastrointestinal bleeding. Haemophilia 13, 224–225 (2007).
81. Cekin, A.H. et al. Blastocystis in patients with gastrointestinal symptoms: A case-control study. BMC
Gastroenterol 12, 122 (2012).
Biology of Foodborne Parasites
82. Petersen, A.M. et al. Active ulcerative colitis is associated with low prevalence of Blastocystis and
Dientamoeba fragilis infection. Scand J Gastroenterol 48, 638–639 (2013). PubMed PMID: 23528075.
83. Willing, B.P. et al. A pyrosequencing study in twins shows that gastrointestinal microbial profiles vary
with inflammatory bowel disease phenotypes. Gastroenterology 139, 1844–1854.e1 (2010).
84. Qin, J. et al. A human gut microbial gene catalogue established by metagenomic sequencing. Nature 464,
59–65 (2010).
85. Schuijt, T.J., van der Poll, T., de Vos, W.M., and Wiersinga, W.J. The intestinal microbiota and host
immune interactions in the critically ill. Trends Microbiol 21, 221–229 (2013).
86. Rajilić-Stojanović, M., Shanahan, F., Guarner, F., and de Vos, W.M. Phylogenetic analysis of dysbiosis in
ulcerative colitis during remission. Inflamm Bowel Dis 19, 481–488 (2013).
87. Joossens, M. et al. Dysbiosis of the faecal microbiota in patients with Crohn’s disease and their unaffected relatives. Gut 60, 631–637 (2011).
88. Vermeiren, J. et al. Decreased colonization of fecal Clostridium coccoides/Eubacterium rectale species from ulcerative colitis patients in an in vitro dynamic gut model with mucin environment. FEMS
Microbiol Ecol 79, 685–696 (2012).
89. Hamad, I., Sokhna, C., Raoult, D., and Bittar, F. Molecular detection of eukaryotes in a single human
stool sample from Senegal. PLoS ONE 7, e40888 (2012).
90. Andersen, L.O., Vedel Nielsen, H., and Stensvold, C.R. Waiting for the human intestinal eukaryotome.
ISME J 7, 1253–1255 (2013). PubMed PMID: 23407309; PubMed Central PMCID: PMC3695289.
91. Scanlan, P.D. Blastocystis: Past pitfalls and future perspectives. Trends Parasitol 28, 327–334 (2012).
92. Poirier, P., Wawrzyniak, I., Vivarès, C.P., Delbac, F., and El Alaoui, H. New insights into Blastocystis
spp.: A potential link with irritable bowel syndrome. PLoS Pathog 8, e1002545 (2012).
93. Arumugam, M. et al. Enterotypes of the human gut microbiome. Nature 473, 174–180 (2011).
94. Rajilić-Stojanović, M. et al. Global and deep molecular analysis of microbiota signatures in fecal samples from patients with irritable bowel syndrome. Gastroenterology 141, 1792–1801 (2011).
95. Yakoob, J. et al. Irritable bowel syndrome: In search of an etiology: Role of Blastocystis hominis. Am J
Trop Med Hyg 70, 383–385 (2004).
96. Yakoob, J. et al. Irritable bowel syndrome: Is it associated with genotypes of Blastocystis hominis.
Parasitol Res 106, 1033–1038 (2010).
97. Giacometti, A., Cirioni, O., Fiorentini, A., Fortuna, M., and Scalise, G. Irritable bowel syndrome in
patients with Blastocystis hominis infection. Eur J Clin Microbiol Infect Dis 18, 436–439 (1999).
98. Jimenez-Gonzalez, D.E. et al. Blastocystis infection is associated with irritable bowel syndrome in a
Mexican patient population. Parasitol Res 110, 1269–1275 (2012).
99. Fouad, S.A., Basyoni, M.M., Fahmy, R.A., and Kobaisi, M.H. The pathogenic role of different Blastocystis
hominis genotypes isolated from patients with irritable bowel syndrome. Arab J Gastroenterol 12, 194–
200 (2011).
100. Elwakil, H.S. and Hewedi, I.H. Pathogenic potential of Blastocystis hominis in laboratory mice. Parasitol
Res 107, 685–689 (2010).
101. Chandramathi, S., Suresh, K., Shuba, S., Mahmood, A., and Kuppusamy, U.R. High levels of oxidative
stress in rats infected with Blastocystis hominis. Parasitology 137, 605–611 (2010).
102. Hussein, E.M., Hussein, A.M., Eida, M.M., and Atwa, M.M. Pathophysiological variability of different
genotypes of human Blastocystis hominis Egyptian isolates in experimentally infected rats. Parasitol Res
102, 853–860 (2008).
103. Bouzid, M., Hunter, P.R., Chalmers, R.M., and Tyler, K.M. Cryptosporidium pathogenicity and virulence. Clin Microbiol Rev 26, 115–134 (2013).
104. Nigro, L. et al. A placebo-controlled treatment trial of Blastocystis hominis infection with metronidazole.
J Travel Med 10, 128–130 (2003).
105. Heyland, K., Friedt, M., Buehr, P., and Braegger, C.P. No advantage for antibiotic treatment over placebo
in Blastocystis hominis-positive children with recurrent abdominal pain. J Pediatr Gastroenterol Nutr 54,
677–679 (2012).
106. Rossignol, J.F., Kabil, S.M., Said, M., Samir, H., and Younis, A.M. Effect of nitazoxanide in persistent diarrhea and enteritis associated with Blastocystis hominis. Clin Gastroenterol Hepatol 3, 987–991
107. Stensvold, C.R., Smith, H.V., Nagel, R., Olsen, K.E., and Traub, R.J. Eradication of Blastocystis carriage
with antimicrobials: Reality or delusion? J Clin Gastroenterol 44, 85–90 (2010).
108. Engsbro, A.L. and Stensvold, C.R. Blastocystis: To treat or not to treat…but how? Clin Infect Dis 55,
1431–1432 (2012).
109. Banaticla, J.E. and Rivera, W.L. Detection and subtype identification of Blastocystis isolates from wastewater samples in the Philippines. J Water Health 9, 128–137 (2011).
110. Lee, L.I., Chye, T.T., Karmacharya, B.M., and Govind, S.K. Blastocystis sp.: Waterborne zoonotic organism, a possibility? Parasit Vectors 5, 130 (2012).
111. Leelayoova, S. et al. Drinking water: A possible source of Blastocystis spp. subtype 1 infection in schoolchildren of a rural community in central Thailand. Am J Trop Med Hyg 79, 401–406 (2008).
112. Leelayoova, S. et al. Evidence of waterborne transmission of Blastocystis hominis. Am J Trop Med Hyg
70, 658–662 (2004).
113. Ithoi, I., Jali, A., Mak, J.W., Wan Sulaiman, W.Y., and Mahmud, R. Occurrence of Blastocystis in water
of two rivers from recreational areas in Malaysia. J Parasitol Res 2011, 123916 (2011).
114. Al-Binali, A.M., Bello, C.S., El-Shewy, K., and Abdulla, S.E. The prevalence of parasites in commonly
used leafy vegetables in South Western, Saudi Arabia. Saudi Med J 27, 613–616 (2006).
115. Turnbaugh, P.J. et al. The human microbiome project. Nature 449, 804–810 (2007).
116. Lozupone, C.A., Stombaugh, J.I., Gordon, J.I., Jansson, J.K., and Knight, R. Diversity, stability and
resilience of the human gut microbiota. Nature 489, 220–230 (2012).
117. Blainey, P.C. The future is now: Single-cell genomics of bacteria and archaea. FEMS Microbiol Rev 37,
407–427 (2013).
118. Yilmaz, S. and Singh, A.K. Single cell genome sequencing. Curr Opin Biotechnol 23, 437–443 (2012).
119. Weinstock, J.V. Autoimmunity: The worm returns. Nature 491, 183–185 (2012).
Lihua Xiao and Una Ryan
Introduction..................................................................................................................................... 77
Morphology and Classification....................................................................................................... 78
Biology, Genetics, and Genomics................................................................................................... 78
5.3.1 Biology................................................................................................................................ 78
5.3.2 Genetics.............................................................................................................................. 80
5.3.3 Genomics............................................................................................................................ 81
5.4 Diagnosis and Typing...................................................................................................................... 82
5.4.1 Microscopy......................................................................................................................... 82
5.4.2 Fecal Antigen Detection..................................................................................................... 83
5.4.3 Genotyping and Subtyping................................................................................................. 83
5.5 Epidemiology and Molecular Epidemiology.................................................................................. 83
5.5.1 Cryptosporidium spp. in Humans...................................................................................... 84
5.5.2 Susceptible Populations...................................................................................................... 84
5.5.3 Anthroponotic versus Zoonotic Transmission................................................................... 84
5.5.4 Waterborne Transmission................................................................................................... 85
5.5.5 Foodborne Transmission.................................................................................................... 86
5.6 Pathogenesis and Clinical Features................................................................................................ 86
5.7 Treatment and Prevention............................................................................................................... 87
Acknowledgments..................................................................................................................................... 87
References................................................................................................................................................. 87
5.1 Introduction
Cryptosporidium spp. inhabit the brush borders of the gastrointestinal, respiratory, and sometimes renal
epithelium of various vertebrates, causing enterocolitis, diarrhea, and cholangiopathy in humans.1,2
Immunocompetent children and adults with cryptosporidiosis usually have a short-term illness accompanied by watery diarrhea, nausea, vomiting, and weight loss. In immunocompromised persons, however, the infection can be protracted and life-threatening. In animals, cryptosporidiosis is an important
cause of diarrhea in neonatal animals. Respiratory and gastric cryptosporidiosis causes substantial mortality in chickens and snakes, respectively.3
The incidence of the disease in humans varies widely among geographic areas. In the United States,
there were 2,769–3,787 annual reported cases during 1999–2002, 3,505–8,269 during 2003–2005,
6,479–11,657 during 2006–2008, and 7,656–8,951 during 2009–2010, corresponding to 1.0–1.3, 1.2–2.8,
2.2–3.9, and 2.5–2.9 cases per 100,000 population, with Midwest states having the highest incidence.4–7
Nevertheless, it is estimated that 98.6% cryptosporidiosis cases remain undiagnosed; thus, there are
about a 3/4 million cases of cryptosporidiosis in the United States each year.8 Cryptosporidiosis exerts
its highest impact in industrialized nations through the occurrence of foodborne, waterborne, person-toperson, and zoonotic outbreaks of diarrhea.9 In developing countries, cryptosporidiosis is a significant
cause of diarrhea and malnutrition in children and AIDS patients.10
Biology of Foodborne Parasites
5.2 Morphology and Classification
Cryptosporidium spp. are obligate protozoan parasites belonging to the family Cryptosporidiidae, which is
a member of the phylum Apicomplexa. The exact placement of Cryptosporidiidae in Apicomplexa is uncertain. It was long considered a member of the class Coccidea, in the order of Eimeriida.3 Recent phylogenetic
and genomic studies, however, indicate that Cryptosporidium spp. are more related to gregarines.11,12
The taxonomy of Cryptosporidium has gone through recent revisions as the result of increased recognition of narrow host specificity by most Cryptosporidium spp. and genetic and biologic characterizations
of parasites from various animals.13,14 Currently, 26 Cryptosporidium species are recognized, including
C. hominis and C. viatorum in humans; C. parvum in ruminants and humans; C. andersoni, C. bovis,
and C. ryanae in cattle and other bovine animals; C. xiaoi in sheep; C. suis and C. scrofarum in pigs;
C. ubiquitum in ruminants, rodents, and primates; C. canis in dogs; C. felis in cats; C. erinacei in horses
and hedgehogs; C. cuniculus in rabbits; C. muris and C. tyzzeri in rodents; C. wrairi in guinea pigs;
C. fayeri and C. macropodum in marsupials; C. meleagridis, C. baileyi, and C. galli in birds; C. ­varanii
and C. serpentis in reptiles; C. fragile in amphibians; and C. molnari in fish. The species status of
Cryptosporidium scophthalmi in fish is yet to be supported by molecular characterization. There are also
many host-adapted Cryptosporidium genotypes that do not yet have species names, such as horse, hamster, ferret, skunk, squirrel, bear, deer, fox, mongoose, panda, wildebeest, seal, duck, woodcock, muskrat
I and II, opossum I and II, chipmunk I–III, rat I–V, deer mouse I–IV, avian I–V, goose I–IV, snake I and
II, tortoise I and II, and piscine I–VIII.13–15 These species/genotypes biologically, morphologically, and
phylogenetically belong to three groups: intestinal, gastric, and piscine species/genotypes.16
The infectious stage of Cryptosporidium is the oocyst (Figure 5.1). This is also the stage shed in the
environment and targeted by most diagnostic assays. Oocysts of intestinal and piscine species are generally spherical and 4–6 μm in size. In contrast, oocysts of gastric species are more elongated and 6–9 μm
in size (Table 5.1). Each oocyst contains four banana-shaped sporozoites. There are no significant morphologic differences among Cryptosporidium species within each group.
5.3 Biology, Genetics, and Genomics
5.3.1 Biology
Cryptosporidium spp. are intracellular parasites that primarily infect epithelial cells of the stomach,
intestine, and biliary ducts. In birds and in severely immunosuppressed persons, the respiratory tract
is sometimes involved. The infection site varies according to species, but almost the entire development of Cryptosporidium spp. occurs between the two lipoprotein layers of the membrane of the
epithelial cells,3 with the exception in C. molnari and other piscine genotypes, for which oogonial
and sporogonial stages are located deep within the epithelial cells.17
FIGURE 5.1 (a) Oocysts of C. hominis under DIC, (b) modified Ziehl–Neelsen acid-fast, and (c) immunofluorescence
Size and Shape Index of Established Cryptosporidium Species
Site of
C. muris
C. parvum
Small intestine
C. wrairi
C. felis
C. andersoni
Cattle, sheep,
goats, horses,
Guinea pigs
C. canis
Small intestine
C. hominis
C. suis
C. bovis
Small intestine
Small and
Small intestine
C. fayeri
C. macropodum
C. ryanae
Small intestine
Small intestine
Small intestine
C. xiaoi
Small intestine
C. ubiquitum
Small intestine
C. cuniculus
Sheep, deer,
C. tyzzeri
C. viatorum
Small intestine
Small intestine
C. scrofarum
Small intestine
C. erinacei
Small intestine
C. meleagridis
C. baileyi
C. galli
C. fragile
C. serpentis
C. varanii
C. molnari
Small intestine
Small intestine
Small intestine
Cloaca, bursa,
Intestine and
Stomach (and
Size (µm)
Shape Index
1.38 (1.25–1.61)
8.4 ± 0.3 × 6.1 ± 0.3 (8.0–9.0 ×
5.6–6.4) (n = 25)
5.0 (4.5–5.4) × 4.5 (4.2–5.0) (n = 50) 1.1 (1.0–1.3)
5.4 (4.8–5.6) × 4.6 (4.0–5.0)
4.6 (3.2–5.1) × 4.0 (3.0–4.0) (n = 40?)
7.4 ± 0.1 × 5.5 ± 0.1 (6.0–8.1 ×
5.0–6.5) (n = 50)
4.95 (3.68–5.88) × 4.71 (3.68–5.88)
(n = 50)
5.2 (4.4–5.9) × 4.9 (4.4–5.4) (n = 100)
4.6 (4.4–4.9) × 4.2 (4.0–4.3) (n = 50)
1.17 (1.04–1.33)
1.1 (1.0–1.2)
1.35 (1.07–1.50)
1.05 (1.04–1.06)
1.07 (1.0–1.09)
4.89 (4.76–5.35) × 4.63 (4.17–4.76)
(n = 50)
4.9 (4.5−5.1) × 4.3 (3.8–5.0) (n = 50)
5.4 (5.0–6.0) × 4.9 (4.5–6.0) (n = 50)
3.73 (2.94–4.41) × 3.16 (2.94–3.68)
(n = 0)
3.94 (2.94–4.41) × 3.44 (2.94–4.41)
(n = 25)
5.04 (4.71–5.32) × 4.66 (4.33–4.98)
(n = 50)
1.14 (1.02–1.18)
5.98 (5.55–6.40) × 5.38 (5.02–5.92)
(n = 50)
4.64 ± 0.05 × 4.19 ± 0.06 (n = 69)
5.35 (4.87–5.87) × 4.72 (4.15–5.20)
(n = 50)
5.16 (4.81–5.96) × 4.83 (4.23–5.29)
(n = 400)
4.9 (4.5–5.8) × 4.4 (4.0–4.8) (n = 100)
1.11 ± 0.02
1.14 (1.03–1.32)
1.07 ± 0.06
1.13 (1.02–1.35)
5.0 (4.5–6.0) × 4.4 (4.2–5.3)
6.2 × 4.6 (5.6–6.3 × 4.5–4.8) (n = 25)
1.1 (1.0–1.3)
1.4 (1.2–1.4)
8.25 (8.0–8.5) × 6.3 (6.2–6.4) (n = 50)
6.2 (5.5–7.0) × 5.5 (5.0–6.5) (n = 50)
5.94 (5.82–6.06) × 5.11 (5.03–5.19)
(n = 37)
4.94 (4.81–5.07) × 4.49 (4.35–4.63)
(n = 20)
4.72 (3.23–5.45) × 4.47 (3.02–5.04)
(n = 22)
1.13 (1.00–1.30)
1.17 (1.14–1.20)
1.14 (1.11–1.17)
1.05 (1.0–1.17)
Biology of Foodborne Parasites
Thick-walled oocyst
ingested by host
Recreational water Drinking water
Contamination of 2
water and
food with oocysts
1 Thick-walled
(sporulated) exist host
Thick-walled oocyst
ingested by host
of environment
with oocysts
1 Thick-walled oocyst
(sporulated) exist host
FIGURE 5.2 Transmission of (a) C. hominis and (b) C. parvum. (Image courtesy of DPDx [
cryptosporidiosis/index.html] from the Centers for Disease Control and Prevention.)
Cryptosporidium infections in humans or other susceptible hosts start with the ingestion of viable
oocysts (Figure 5.2). Upon contact with gastric and duodenal fluid, four sporozoites are liberated from
each excysted oocyst, invade the epithelial cells, and develop to trophozoites surrounded by a parasitophorous vacuole. Within the epithelial cells, trophozoites undergo 2–3 generations of asexual amplification in a process called merogony, leading to the formation of different types of meronts containing
4–8 merozites. The latter differentiate into sexually distinct stages called macrogamonts and microgamonts (containing microgametes) in the process of gametogony. Zygotes are formed in the epithelial cells
from the fusion of macrogamonts and microgametes, developing into oocysts, which sporulate in situ in
the process of sporogony, and contain four sporozoites. It is believed that about 20% of oocysts are thin
walled and may excyst within the digestive tract of the host, leading to autoinfection. The thick-walled
oocysts are excreted into the environment; are resistant to low temperature, high salinity, and most
disinfectants; and can initiate infection in a new host upon ingestion. The prepatent period (the time
from ingestion of infective oocysts to the completion of endogenous development and excretion of new
oocysts) varies with species, hosts, and infection doses; it is usually between 4 and 10 days for intestinal species.3,18 In addition to the classic coccidian developmental stages, a gregarine-like extracellular
stage was recently described in C. andersoni and C. parvum, which might go through multiplication
via syzygy, a sexual reproduction process involving the end-to-end fusion of two or more parasites.19,20
5.3.2 Genetics
Cryptosporidium spp. are haploid organisms. Because of the existence of a sexual phase in its life cycle,
genetic recombination is possible between two related subtypes. Thus, new subtypes can emerge as a
result of sexual recombination during concurrent infection of mixed populations, which was demonstrated by experimental infection of mice with different subtypes of C. parvum.21 Indeed, multilocus
characterizations of field isolates have mostly identified a panmictic population structure of C. parvum
in cattle in many areas.22–24 However, in some areas such as Italy, a clonal genetic population structure in
C. parvum in cattle has been shown.25 The population structure of Cryptosporidium spp. is likely influenced by the intensity of transmission, the host–parasite interaction, and management practices. Rather
than conforming to a strict paradigm of either a clonal, panmictic, or epidemic population structure, a
flexible reproductive strategy characterized by the co-occurrence of these mechanisms is frequently used
by C. parvum. The relative contribution of each pathway appears to vary between the regions, perhaps
dependent on the prevailing ecological determinants of transmission.26–28
Host adaptation has been seen in C. parvum, which infects several species of mammals, indicating the
occurrence of genetic isolation within some parasite lineages. The presence of host-adapted C. parvum
subtype families is well known at the 60 kDa glycoprotein (gp60, also known as gp15/45/60, gp40/15)
locus, including IIa in cattle, IId in sheep and goats, and IIc and IIe in humans.29 The existence of these
host-adapted C. parvum subpopulations has been confirmed by multilocus characterization of parasites in
several European countries.24,25,30,31 Host adaptation has recently been shown also to occur in C. ubiquitum.32
In contrast to the diversity in population structure of C. parvum, C. hominis usually maintains a
clonal population structure.24,33,34 In some areas such as Europe, there is only limited heterogeneity
in C. ­hominis compared with C. parvum. This appears to be due to an expansion of a single subtype
(the gp60 subtype IbA10G2),24 as would be the case in an epidemic clonal structure. It does not appear
this is the case in high endemic areas such as India.34 Geographically segregated subpopulations of
C. ­hominis are often seen in developing countries where intercountry population exchange is limited.33,35
No s­ ignificant segregation in parasite population is seen in C. parvum in animals.23 Despite an overall
clonal population genetic structure in C. hominis, it has been shown recently that genetic recombination
is a driving force in the emergence of virulent subtypes.36,37 Genetic recombination is also responsible for
the success of the dominant C. parvum IIaA15G2R1 subtype.38
There is no established toolbox for genetic manipulation of Cryptosporidium spp.39 Genetic crossing
between two C. parvum isolates is feasible in a laboratory animal model,21 but there are no facile techniques that can be used to screen for recombinant progenies. Transient transfection of C. parvum oocysts
with a dsRNA virus vector containing the green fluorescent protein as marker was reported.40 However,
there are currently no drug resistance markers and easily typeable phenotypes for selection of progenies.
The lack of genetic manipulation tools has greatly hampered our understanding of the virulence and
invasion mechanisms of Cryptosporidium spp.
5.3.3 Genomics
The genome of three Cryptosporidium species has been sequenced using the traditional Sanger sequencing
technology, including C. parvum, C. hominis, and C. muris.41,42 ( More recently,
an isolate each of the host-adapted C. parvum IIc subtype family and C. ubiquitum has been sequenced using
the next-generation sequencing technology.32,43 The C. parvum genome is fairly complete, with 21 ­contigs
and only 10 physical gaps distributed among 8 chromosomes. The 9.1 Mb genome has a 30% G+C content and is much smaller than those of Plasmodium falciparum (22.8 Mb), Toxoplasma gondii (63 Mb), and
Ascogregarina taiwanensis (~30 Mb) but slightly larger than those of Theileria spp. (8.3 Mb) (Table 5.2).
Major Features of Cryptosporidium Genome in Comparison with Other Apicomplexans
Feature or Pathway
C. parvum
T. gondii
P. falciparum
T. parva
A. taiwanensis
Genome sizes (Mb)
G+C contents (%)
Protein-encoding genes
Apicoplast genome (kb)
Mitochondrial genome
De novo synthesis
Amino acids
De novo
De novo
De novo
De novo
Fatty acid synthesis
Type II (plastid, de novo)
Type I (FA elongation)
Polyketide synthase
Mannitol cycle
Ethanol production
Sources: Based on Templeton, T.J. et al., Mol. Biol. Evol., 27, 235, 2010; Zhu, G. and Xiao, L., Cryptosporidium
species, in Fratamico, P., Liu, Y., and Kathariou, S., eds., Genomes of Foodborne and Waterborne
Pathogens, American Society for Microbiology, Washington, DC, 2011, pp. 271–286.
Biology of Foodborne Parasites
The compact C. parvum genome is largely a result of the loss of many metabolic pathways, such as the
Krebs cycle, cytochrome-based respiratory chain, and major de novo synthetic pathways for amino acids,
nucleotides, and fatty acids, as well as the small intergenic regions and the rarity of introns (the only gene with
confirmed introns is the β-tubulin gene).41 Additionally, the parasite has only a single copy of almost all genes
except for rRNA genes and lacks large gene families such as var genes in P. falciparum. Unlike most other
apicomplexans, it also has no apicoplast and mitochondrial genomes. Because of the lack of de novo synthesis
capacities of amino acids, nucleotides, and fatty acids, C. parvum has an array of expanded families of transporters to acquire nutrients from the host, including at least 11 transporters for amino acids, 20 for sugars, and
19 annotated ATP-binding cassettes (ABCs) for transporting various metabolites, lipids/sterols, and drugs.
These transporters, especially ABC transporters, are probably under selection pressure, as they are among the
most polymorphic genes in the C. parvum genome.43
The published C. hominis genome is fragmented and contains 1413 contigs with ~246 gaps. It has an
almost perfect synteny with that of C. parvum, as the average identities at their nucleotide and amino
acid levels are about 97% and 98%, respectively. Therefore, the two genomes encode almost identical sets
of genes. There is a difference in the number of annotated genes between C. parvum and C. hominis,41,42
but this is largely due to the use of different strategies and stringencies in gene prediction. The sequenced
C. muris genome is also fragmental, although less than that of C. hominis. It has a much lower synteny
and sequence similarity to genomes of C. parvum and C. hominis. Thus, there are significant gene insertions and deletions in the C. muris genome. For example, the C. muris genome has the full components
for the Krebs cycle and a functional ATP synthase.44 Therefore, the mitosome of C. muris functions
essentially as a typical mitochondrion. In contrast, C. parvum and C. hominis rely on glycolysis and
substrate-level phosphorylation in the mitosome for the production of ATP.
The complete genome sequences and annotations for C. parvum, C. hominis, and C. muris are available at the Cryptosporidium genome database CryptoDB (, which is a member of the
EukPathDB (
5.4 Diagnosis and Typing
At the moment, almost all active Cryptosporidium infections are diagnosed by analysis of stool specimens. Stool specimens are usually collected fresh or in fixative solutions such as 10% buffered formalin and polyvinyl alcohol (PVA).45 However, stool specimens fixed in formalin and mercury-based
preservatives (such as LV-PVA) cannot be used for PCR, which requires the use of fresh or frozen stool
specimens or stools preserved in TotalFix, zinc PVA, or 2.5% potassium dichromate. In clinical laboratories, Cryptosporidium spp. in stool specimens are commonly detected by microscopic examinations of
oocysts or immunologic detection of antigens.46 PCR-based typing methods, however, are increasingly
used in investigations of outbreaks, surveillance, and diagnosis of cryptosporidiosis.29
5.4.1 Microscopy
Stool specimens can be examined for Cryptosporidium oocysts by microscopy of direct wet mount,
after oocyst concentration by traditional ethyl acetate or Weber-modified ethyl acetate concentration
methods.46 Bright-field or differential interference contrast (DIC) microscopy of direct wet mounts is
generally used. This allows the observation of oocyst morphology and more accurate measurement of
oocysts (Figure 5.1). Most Cryptosporidium species look similar under microscopes and have similar
morphometric measurements, although Cryptosporidium oocysts in humans are generally 4–6 μm in
size13 (Table 5.1).
More often, Cryptosporidium oocysts in concentrated stool specimens are detected by microscopy after
staining of the fecal smears. Many special stains have been used in the detection of Cryptosporidium
oocysts, but modified acid-fast stains are the most commonly used,46 especially in developing countries,
because of their low cost and simultaneous detection of several other pathogens such as Cystoisospora and
Cyclospora. Two stains widely used are the modified Ziehl–Neelsen acid-fast stain and modified Kinyoun’s
acid-fast stain.46 Oocysts are stained bright red to purple against blue or green background (Figure 5.1).
Direct immunofluorescence assays (DFA) are used increasingly in Cryptosporidium oocyst detection, especially in industrialized nations. Compared to acid-fast staining, DFA has higher sensitivity and
specificity.47 Many commercial DFA kits are marketed for the diagnosis of Cryptosporidium, most of
which include reagents allowing simultaneous detection of Giardia cysts. Oocysts appear apple green
against a dark background in immunofluorescence microscopy (Figure 5.1). It has been shown that most
antibodies in commercial DFA kits react with oocysts of almost all Cryptosporidium species, making
species diagnosis impossible.48,49
5.4.2 Fecal Antigen Detection
Cryptosporidium infection can also be diagnosed by the detection of Cryptosporidium antigens in stool
specimens by immunoassays.46 Antigen-capture-based enzyme immunoassays (EIA) have been used
in the diagnosis of cryptosporidiosis since 1990. In recent years, EIAs have gained popularity because
they do not require experienced microscopists and can be used to screen a large number of samples.50
Several commercial EIA kits are commonly used in clinical laboratories. High specificity (99%–100%)
has been generally reported for these EIA kits.47,51,52 Sensitivities, however, have been reported to range
from 70%47 to 94%–100%.51,53–55 Most EIA kits have been evaluated only with human stool specimens
presumably from patients infected with C. hominis or C. parvum.55 Their usefulness in the detection of
Cryptosporidium spp. in animals may be compromised by the specificity of the antibodies.
In recent years, lateral-flow immunochromatographic assays have gained popularity in rapid detection
of Cryptosporidium in stool specimens. In evaluation studies, these assays have been shown to have high
specificities (>90%) and sensitivity (98%–100%).47,56–61 However, sensitivities of 68%–75% were shown
in some studies for some assays.47,62–64 High false-positive rates (positive predictive value = 56%) of
several rapid assays have been recently reported in clinical diagnosis of cryptosporidiosis in the Unites
States.65 This has prompted the Council of State and Territorial Epidemiologists to change the case
definition of rapid assay-positive cases from confirmed cases to probable cases. It has also been shown
recently that some rapid assays kits have low sensitivity (<35%) in detecting some Cryptosporidium species other than C. hominis and C. parvum.64
5.4.3 Genotyping and Subtyping
Molecular techniques, especially PCR and PCR-related methods, have been developed and used in the
detection and differentiation of Cryptosporidium spp. for many years. Several genus-specific PCR–RFLPbased genotyping tools have been developed for the detection and differentiation of Cryptosporidium
at the species level.66–70 Most of these techniques are based on the small subunit (SSU) rRNA gene.
Other genotyping techniques are designed mostly for the differentiation of C. parvum and C. hominis,
thus cannot detect and differentiate other Cryptosporidium spp. or genotypes.29 Their usefulness in the
analysis of human stool specimens is compromised by their inability to detect C. canis, C. felis, C. suis,
C. muris, and other species/genotypes divergent from C. parvum and C. hominis.71
Several subtyping tools have been developed to characterize the diversity within C. parvum or C. hominis.29 One of the most commonly used techniques is DNA sequence analysis of the gp60 gene.72–75 Most
of the genetic heterogeneity in this gene is present in the number of a trinucleotide repeat (TCA, TCG,
or TCT), although extensive sequence differences in nonrepeat regions are also present between subtype
families. Multilocus subtyping tools for C. parvum and C. hominis have also been developed.24,26,31,76,77
The usefulness of subtyping tools has been demonstrated by the analysis of samples from foodborne and
waterborne outbreaks of cryptosporidiosis.30,78–92
5.5 Epidemiology and Molecular Epidemiology
Humans can acquire cryptosporidiosis through several transmission routes, such as direct contact with
infected persons or animals and consumption of contaminated water (drinking or recreational) or food.1
The transmission routes differ between the two dominant species in humans, C. parvum and C. hominis
Biology of Foodborne Parasites
(Figure 5.2). The relative role of each transmission route in the epidemiology of cryptosporidiosis and
the infection sources are frequently unclear for a particular country or area.
5.5.1 Cryptosporidium spp. in Humans
Currently, nearly 20 Cryptosporidium species and genotypes have been reported in humans, including
C. hominis, C. parvum, C. meleagridis, C. felis, C. canis, C. ubiquitum, C. ­cuniculus, C. ­viatorum,
C. muris, C. suis, C. andersoni, C. tyzzeri, C. fayeri, C. bovis, C. scrofarum, C. ­erinacei, and
Cryptosporidium horse, skunk genotypes and chipmunk genotype I. Humans are most frequently
infected with C. hominis and C. parvum. The former infects almost exclusively humans and ­nonhuman
primates, and is thus considered an anthroponotic parasite, whereas the latter mostly infects humans,
ruminants, and horses, and thus considered a potential zoonotic pathogen. Other species such as
C. meleagridis, C. felis, C. canis, C. ubiquitum, C. cuniculus, and C. viatorum are less common.
The remaining species and genotypes have been only found in a few human cases.29,93–97 These
Cryptosporidium spp. infect both immunocompetent and immunocompromised persons. The distribution of these species in humans is different among geographic areas and socioeconomic conditions,
with C. canis and C. felis mostly seen in humans in developing countries, C. ubiquitum mostly in
industrialized nations, and C. cuniculus mostly in the United Kingdom. This is probably the result of
differences in infection sources and transmission routes.29
5.5.2 Susceptible Populations
In developing countries, human Cryptosporidium infection occurs mostly in children younger than
2 years.10,18 In developed countries, pediatric cryptosporidiosis occurs in children later than in developing countries, probably due to delayed exposures to contaminated environments as a result of better hygiene.4,6,7,73,95 Cryptosporidiosis is also common in elderly people attending nursing homes where
person-to-person transmission probably plays a major role in the spread of Cryptosporidium infections.98
In the general population, a substantial number of adults are susceptible to Cryptosporidium infection,
as sporadic infections occur in all age groups in the United States and United Kingdom, and traveling
to developing countries and consumption of contaminated food or water can frequently lead to infection.9,99–101 Cryptosporidiosis is common in immunocompromised persons, including AIDS patients, persons with primary immunodeficiency, and cancer and transplant patients undergoing immunosuppressive
therapy.1,102–105 Hemodialysis patients with chronic renal failure and renal transplant patients commonly
develop cryptosporidiosis.105–109 In HIV+ persons, the occurrence of cryptosporidiosis increases as the
CD4+ lymphocyte cell counts fall, especially below 200 cells/μL.103
5.5.3 Anthroponotic versus Zoonotic Transmission
In the United States and Europe, contact with persons with diarrhea has been identified as a major risk
factor for sporadic cryptosporidiosis.87,100,110–112 The importance of direct person-to-person transmission
is exemplified by the high prevalence of cryptosporidiosis in childcare facilities, nursing homes, and
mothers with young children in these countries. In most of these case–control studies, contact with farm
animals (especially cattle) is also a major risk factor for human cryptosporidiosis.9,100,110,111,113,114
Differences in the distribution of Cryptosporidium genotypes in humans are a reflection of differences in infection sources.29,114–116 The occurrence of C. hominis in humans is most likely due to anthroponotic transmission, whereas C. parvum in a population can be the result of both anthroponotic and
zoonotic transmission. Thus far, studies conducted in developing countries have shown a predominance
of C. hominis in children or HIV+ adults. This is also true for most areas in the United States, Canada,
Australia, and Japan. In Europe and New Zealand, however, several studies have shown almost equal
prevalence of C. parvum and C. hominis in both immunocompetent and immunocompromised persons.29 Thus, in most developing countries, anthroponotic transmission of Cryptosporidium plays a
major role in human cryptosporidiosis, whereas in Europe, New Zealand, and rural areas of the United
States, both anthroponotic and zoonotic transmissions are important. In Mideast countries, children are
mostly infected with C. parvum, but the significance of this is not clear.117
Sequences analyses of the gp60 gene have shown that many C. parvum infections in humans are
not the result of zoonotic transmission.29 Among several C. parvum subtype families identified, IIa
and IIc are the two most common families. The former has been identified in both humans and calves,
thus can be a zoonotic pathogen, whereas IIc has only been seen in humans,16,29,74 thus an anthroponotic pathogen. In developing countries, most C. parvum infections in children and HIV+ persons
are caused by the subtype family IIc, with IIa largely absent, indicating that anthroponotic transmission of C. parvum is common in these areas.16,29 In contrast, both IIa and IIc subtype families are
seen in humans in developed countries, even in the United Kingdom, where zoonotic transmission is
known to play a significant role in the transmission of human cryptosporidiosis.118 Another C. parvum
subtype family commonly found in sheep and goats, IId, is the dominant C. parvum subtype family
in humans in Mideast countries.117 Results of multilocus subtyping support the conclusions of gp60
subtyping studies.31,119
5.5.4 Waterborne Transmission
Epidemiologic studies have frequently identified water as a major route of Cryptosporidium transmission
in industrialized nations. Numerous outbreaks of cryptosporidiosis have been associated with drinking
water in these countries.120 Seasonal variations in the incidence of human Cryptosporidium infection in
these countries have been partially attributed to waterborne transmission.9,99,100,121 Thus, in the United
States, there is a late summer peak in sporadic cases of cryptosporidiosis,7,9,100 which is largely due to
recreational activities such as swimming and water sports.122 In the United States and Canada, swimming in a lake or river was identified as a risk factor.100,112
The role of drinking water in sporadic Cryptosporidium infection is not clear. In England, there is
an association between numbers of glasses of tap water drunk at home each day and the occurrence
of sporadic cryptosporidiosis.110 In the United States, drinking untreated surface water was identified
as a risk factor for the acquisition of Cryptosporidium in a small case–control study,123 and residents
living in cities with surface-derived drinking water generally have higher blood antibody levels against
Cryptosporidium than those living in cities with groundwater as drinking water, indicating that drinking
water plays a role in the transmission of human cryptosporidiosis.124 Nevertheless, case–control studies
conducted in the United States and Canada have failed to show a direct linkage of Cryptosporidium
infection to drinking water.125–127
Numerous waterborne outbreaks of cryptosporidiosis have occurred in the United States, Canada,
United Kingdom, France, Australia, Japan, and other industrialized nations.120,128–130 These include
outbreaks associated with both drinking water and recreational water (swimming pools and water
parks). After the massive cryptosporidiosis outbreak in Milwaukee in 1993, several new drinking water
regulations have been implemented in the United States, United Kingdom, and other industrialized
nations.131–133 As a result of more stringent treatment of source water, the number of drinking water–­
associated outbreaks is in decline in these countries, and most recent outbreaks in the United States
are associated with recreational water.9,122 Even though five Cryptosporidium species are commonly
found in humans, C. parvum and C. hominis are responsible for most cryptosporidiosis outbreaks,
with C. ­hominis responsible for more outbreaks than C. parvum.29 This is even the case for the United
Kingdom, where C. ­parvum and C. hominis are both common in the general population. Recently, there
was one drinking water–­associated cryptosporidiosis outbreak caused by C. cuniculus.134
The importance of waterborne transmission in cryptosporidiosis epidemiology is not clear in developing countries. In most tropical countries, Cryptosporidium infections in children usually peak during the
rainy season; thus, waterborne transmission probably plays a role in the transmission of cryptosporidiosis in these areas.135–137 In Mexican children living near the US border, cryptosporidiosis is associated
with consumption of municipal water instead of bottled water.138
Biology of Foodborne Parasites
5.5.5 Foodborne Transmission
The role of food in the transmission of cryptosporidiosis is much less clear. Cryptosporidium oocysts
have been isolated from several foodstuffs, and these have mainly been associated with fruits, vegetables, and shellfish.139–141 Direct contamination of food by fecal materials from animals or food handlers
has been implicated in several foodborne outbreaks of cryptosporidiosis in industrialized nations. In
most instances, human infections were acquired through consumption of contaminated fresh produce
and unpasteurized apple cider or milk.82,142–146
Very few case–control studies have examined the role of contaminated food as a risk factor in the
acquisition of Cryptosporidium infection in endemic areas. A pediatric study in Brazil failed to show
any association between Cryptosporidium infection and diet or type of food hygiene.147 Case–control
studies conducted in the United States, United Kingdom, and Australia have actually shown a lower
prevalence of Cryptosporidium infection in immunocompetent persons with frequent consumption of
raw vegetables.87,100,110,148 Nevertheless, it is estimated that about 8% of Cryptosporidium infections in
the United States are foodborne.8
5.6 Pathogenesis and Clinical Features
In developing countries, frequent symptoms of cryptosporidiosis in children include diarrhea, abdominal
cramps, nausea, vomiting, headache, fatigue, and low-grade fever. The diarrhea can be voluminous and
watery but usually resolves within 1–2 weeks without treatment. Not all infected children have diarrhea
or other gastrointestinal symptoms, and the occurrence of diarrhea in children with cryptosporidiosis
can be as low as 30% in community-based studies.149 Even subclinical cryptosporidiosis exerts a significant adverse effect on child growth, as infected children with no clinical symptoms experience growth
faltering, both in weight and in height.1 Children can have multiple episodes of cryptosporidiosis; thus,
immunity against cryptosporidiosis in children is short lived or incomplete.150 Cryptosporidiosis has
been associated with increased mortality in hospitalized children in developing countries.135,151 There
are significant differences among different Cryptosporidium species and C. hominis subtype families in
clinical manifestations of pediatric cryptosporidiosis.150
Immunocompetent persons with sporadic cryptosporidiosis in industrialized nations usually
develop diarrhea.1,101,148 The median number of stools per day during the worst period of the infection is 7–9.5.148 Other common symptoms include abdominal pain, nausea, vomiting, and low-grade
fever.101,148 The duration of illness has a mean or median of 9–21 days, with a median of 5 lost work
or study days and hospitalization of 7%–22% of patients.99,101,148 Patients infected with C. hominis are
more likely to have joint pain, eye pains, recurrent headache, dizziness, and fatigue than those infected
with C. parvum.152
Cryptosporidiosis in immunocompromised persons, including AIDS patients, is frequently associated
with chronic, life-threatening diarrhea.103 Sclerosing cholangitis and other biliary involvements are also
common in AIDS patients with cryptosporidiosis.153 Cryptosporidiosis in AIDS patients is associated
with increased mortality and shortened survival.154 Variations in the infection site (gastric infection,
proximal small intestine infection, ileocolonic infection, vs. pan-enteric infection) have been seen in
AIDS patients with cryptosporidiosis. They may contribute to differences in clinical presentations and
survival.155,156 Likewise, different Cryptosporidium species and C. hominis subtype families are associated with different clinical manifestations in HIV+ persons in developing countries.157
Cryptosporidium spp. commonly infect the colonic and ileal mucosa. However, developing stages may
be found in the entire gastrointestinal tract and part of the respiratory tract in immunocompromised persons. Histologically, mononuclear cell infiltration in the lamina propria, mucosal cell apoptosis, mucosal
inflammation with villus blunting, and cryptitis are usually seen, leading to the loss of barrier function
and malabsorption. Infection of the crypts usually leads to more severe disease, as does infection of
the proximal small bowel with villus flattening.1,153,155,158 The voluminous diarrhea is accompanied with
chloride secretion and impaired glucose absorption and is probably mediated by substance P, a gastrointestinal neuropeptide.159 CD4+ T lymphocytes and Th-1 immune responses play a key role in acquired
immunity against cryptosporidiosis, although CD8+ T lymphocytes contribute to the clearance of the
parasite.158,160 These protective immune responses appear to be mediated through TLR4/NF-kappaBdependent nitric oxide production.161,162
5.7 Treatment and Prevention
Numerous pharmaceutical compounds have been screened for anti-Cryptosporidium activities in vitro
or in laboratory animals.163–165 Some of those showing promises have been used in the experimental
treatment of cryptosporidiosis in humans, but few have been proven effective in controlled clinical trials.103,166 Oral or intravenous rehydration and antimotility drugs are widely used in treating the severe
diarrhea associated with cryptosporidiosis. Nitazoxanide (NTZ) is the only FDA-approved drug for the
treatment of cryptosporidiosis in immunocompetent persons. Clinical trials have demonstrated that NTZ
can shorten clinical disease and reduce parasite loads.164,167 This drug, however, is not effective in the
treatment of Cryptosporidium infections in immunodeficient patients.166,167
In industrialized nations, the most effective treatment and prophylaxis for cryptosporidiosis in AIDS
patients is the use of highly active antiretroviral therapy (HAART).164,165,168 It is probably also an effective prevention for cryptosporidiosis in HIV+ persons in developing countries.169 It is believed that the
eradication and prevention of the infection are related to the replenishment of CD4+ cells in treated
persons and the antiparasitic activities of the protease inhibitors used in HAART.165,168 Relapse of cryptosporidiosis is common in AIDS patients who have stopped taking HAART.170 In developing countries,
protease inhibitors are generally not included in the HAART regimens. Limited reports have shown that
cryptosporidiosis is still common in HIV-positive patients receiving HAART in developing countries,
although at lower frequencies than those generally reported in untreated HIV patients.169,171,172
Good hygiene is the key in preventing the acquisition of Cryptosporidium infection.173
Immunosuppressed persons especially should take necessary precautions to prevent the occurrence of
cryptosporidiosis.174 This includes washing hands before eating and after going to the bathroom, changing diapers, and contacting pets or soil (including gardening); avoiding drinking water from lakes and
rivers, swallowing water in recreational activities, and unpasteurized milk, milk products, and juices;
and following safe-sex practices (avoiding oral–anal contact). During cryptosporidiosis outbreaks or
when a community advisory to boil water is issued, individuals should boil water for 1 min to kill the
parasite or use a tap water filter capable of removing particles <1 μm in diameter. Immunosuppressed
persons also should avoid eating raw shellfish and should not eat uncooked vegetables and unpeeled
fruits when traveling to developing countries.174
The findings and conclusions in this report are those of the authors and do not necessarily represent the
views of the Centers for Disease Control and Prevention.
1. Chalmers, R.M. and Davies, A.P. Minireview: Clinical cryptosporidiosis. Exp Parasitol 124, 138–146
2. Kotloff, K.L. et al. Burden and aetiology of diarrhoeal disease in infants and young children in developing countries (the Global Enteric Multicenter Study, GEMS): A prospective, case-control study. Lancet
382, 209–222 (2013).
3. Fayer, R. Introduction. In Cryptosporidium and Cryptosporidiosis, 2nd edn. (Fayer, R. and Xiao, L.,
eds.), pp. 1–42 (Taylor & Francis Group, Boca Raton, FL, 2008).
4. Yoder, J.S. and Beach, M.J. Cryptosporidiosis surveillance—United States, 2003–2005. MMWR Surveill
Summ 56, 1–10 (2007).
5. Hlavsa, M.C., Watson, J.C., and Beach, M.J. Cryptosporidiosis surveillance—United States 1999–2002.
MMWR Surveill Summ 54, 1–8 (2005).
Biology of Foodborne Parasites
6. Yoder, J.S., Harral, C., and Beach, M.J. Cryptosporidiosis surveillance—United States, 2006–2008.
MMWR Surveill Summ 59, 1–14 (2010).
7. Yoder, J.S., Wallace, R.M., Collier, S.A., Beach, M.J., and Hlavsa, M.C. Cryptosporidiosis s­ urveillance—
United States, 2009–2010. MMWR Surveill Summ 61, 1–12 (2012).
8. Scallan, E. et al. Foodborne illness acquired in the United States—Major pathogens. Emerg Infect Dis
17, 7–15 (2011).
9. Yoder, J.S. and Beach, M.J. Cryptosporidium surveillance and risk factors in the United States. Exp
Parasitol 124, 31–39 (2010).
10. Mor, S.M. and Tzipori, S. Cryptosporidiosis in children in sub-Saharan Africa: A lingering challenge.
Clin Infect Dis 47, 915–921 (2008).
11. Leander, B.S. Marine gregarines: Evolutionary prelude to the apicomplexan radiation? Trends Parasitol
24, 60–67 (2008).
12. Templeton, T.J. et al. A genome-sequence survey for Ascogregarina taiwanensis supports evolutionary
affiliation but metabolic diversity between a gregarine and Cryptosporidium. Mol Biol Evol 27, 235–248
13. Xiao, L., Fayer, R., Ryan, U., and Upton, S.J. Cryptosporidium taxonomy: Recent advances and implications for public health. Clin Microbiol Rev 17, 72–97 (2004).
14. Fayer, R. Taxonomy and species delimitation in Cryptosporidium. Exp Parasitol 124, 90–97 (2010).
15. Kvac, M. et al. Cryptosporidium erinacei n. sp. (Apicomplexa: Cryptosporidiidae) in hedgehogs. Vet
Parasitol 201, 9–17 (2014).
16. Xiao, L. and Feng, Y. Zoonotic cryptosporidiosis. FEMS Immunol Med Microbiol 52, 309–323 (2008).
17. Alvarez-Pellitero, P. et al. Cryptosporidium scophthalmi n. sp. (Apicomplexa: Cryptosporidiidae) from
cultured turbot Scophthalmus maximus. Light and electron microscope description and histopathological
study. Dis Aquat Organ 62, 133–145 (2004).
18. Leitch, G.J. and He, Q. Cryptosporidiosis—An overview. J Biomed Res 25, 1–16 (2012).
19. Rosales, M.J., Cordon, G.P., Moreno, M.S., Sanchez, C.M., and Mascaro, C. Extracellular like-gregarine
stages of Cryptosporidium parvum. Acta Trop 95, 74–78 (2005).
20. Hijjawi, N.S., Meloni, B.P., Ryan, U.M., Olson, M.E., and Thompson, R.C. Successful in vitro cultivation of Cryptosporidium andersoni: Evidence for the existence of novel extracellular stages in the life
cycle and implications for the classification of Cryptosporidium. Int J Parasitol 32, 1719–1726 (2002).
21. Feng, X., Rich, S.M., Tzipori, S., and Widmer, G. Experimental evidence for genetic recombination in
the opportunistic pathogen Cryptosporidium parvum. Mol Biochem Parasitol 119, 55–62 (2002).
22. De Waele, V. et al. Panmictic structure of the Cryptosporidium parvum population in irish calves:
Influence of prevalence and host movement. Appl Environ Microbiol 79, 2534–2541 (2013).
23. Herges, G.R. et al. Evidence that Cryptosporidium parvum populations are panmictic and unstructured in
the Upper Midwest of the United States. Appl Environ Microbiol 78, 8096–8101 (2012).
24. Mallon, M. et al. Population structures and the role of genetic exchange in the zoonotic pathogen
Cryptosporidium parvum. J Mol Evol 56, 407–417 (2003).
25. Drumo, R. et al. Evidence of host-associated populations of Cryptosporidium parvum in Italy. Appl
Environ Microbiol 78, 3523–3529 (2012).
26. Tanriverdi, S. et al. Inferences about the global population structure of Cryptosporidium parvum and
Cryptosporidium hominis. Appl Environ Microbiol 74, 7227–7234 (2008).
27. Ngouanesavanh, T. et al. Cryptosporidium population genetics: Evidence of clonality in isolates from
France and Haiti. J Eukaryot Microbiol 53, S33–S36 (2006).
28. Morrison, L.J. et al. The population structure of the Cryptosporidium parvum population in Scotland: A
complex picture. Infect Genet Evol 8, 121–129 (2008).
29. Xiao, L. Molecular epidemiology of cryptosporidiosis: An update. Exp Parasitol 124, 80–89 (2010).
30. Leoni, F., Mallon, M.E., Smith, H.V., Tait, A., and McLauchlin, J. Multilocus analysis of Cryptosporidium
hominis and Cryptosporidium parvum from sporadic and outbreak-related human cases and C. parvum
from sporadic cases in livestock in the UK. J Clin Microbiol 45, 3286–3294 (2007).
31. Grinberg, A. et al. Host-shaped segregation of the Cryptosporidium parvum multilocus genotype repertoire. Epidemiol Infect 136, 273–278 (2008).
32. Li, N. et al. Subtyping Cryptosporidium ubiquitum, a zoonotic pathogen emerging in humans. Emerg
Infect Dis 20, 217–224 (2014).
33. Gatei, W. et al. Unique Cryptosporidium population in HIV-infected persons, Jamaica. Emerg Infect Dis
14, 841–843 (2008).
34. Gatei, W. et al. Multilocus sequence typing and genetic structure of Cryptosporidium hominis from children in Kolkata, India. Infect Genet Evol 7, 197–205 (2007).
35. Gatei, W. et al. Molecular analysis of the 18S rRNA gene of Cryptosporidium parasites from patients
with or without human immunodeficiency virus infections living in Kenya, Malawi, Brazil, the United
Kingdom, and Vietnam. J Clin Microbiol 41, 1458–1462 (2003).
36. Li, N. et al. Genetic recombination and Cryptosporidium hominis virulent subtype IbA10G2. Emerg
Infect Dis 19, 1573–1582 (2013).
37. Feng, Y., Tiao, N., Li, N., Hlavsa, M., and Xiao, L. Multilocus sequence typing of an emerging
Cryptosporidium hominis subtype in the United States. J Clin Microbiol 52, 524–530 (2014).
38. Feng, Y. et al. Population genetic characterisation of dominant Cryptosporidium parvum subtype
IIaA15G2R1. Int J Parasitol 43, 1141–1147 (2013).
39. Striepen, B. Parasitic infections: Time to tackle cryptosporidiosis. Nature 503, 189–191 (2013).
40. Li, W. et al. Transient transfection of Cryptosporidium parvum using green fluorescent protein (GFP) as
a marker. Mol Biochem Parasitol 168, 143–148 (2009).
41. Abrahamsen, M.S. et al. Complete genome sequence of the apicomplexan, Cryptosporidium parvum.
Science 304, 441–445 (2004).
42. Xu, P. et al. The genome of Cryptosporidium hominis. Nature 431, 1107–1112 (2004).
43. Widmer, G. et al. Comparative genome analysis of two Cryptosporidium parvum isolates with different
host range. Infect Genet Evol 12, 1213–1221 (2012).
44. Mogi, T. and Kita, K. Diversity in mitochondrial metabolic pathways in parasitic protists Plasmodium
and Cryptosporidium. Parasitol Int 59, 305–312 (2010).
45. Garcia, L.S., Bruckner, D.A., Brewer, T.C., and Shimizu, R.Y. Techniques for the recovery and identification of Cryptosporidium oocysts from stool specimens. J Clin Microbiol 18, 185–190 (1983).
46. Smith, H.V. Diagnostics. In Cryptosporidium and Cryptosporidiosis, 2nd edn. (Fayer, R. and Xiao, L.,
eds.), pp. 173–207 (CRC Press, Boca Raton, FL, 2008).
47. Johnston, S.P., Ballard, M.M., Beach, M.J., Causer, L., and Wilkins, P.P. Evaluation of three commercial
assays for detection of Giardia and Cryptosporidium organisms in fecal specimens. J Clin Microbiol
41, 623–626 (2003).
48. Graczyk, T.K., Cranfield, M.R., and Fayer, R. Evaluation of commercial enzyme immunoassay (EIA)
and immunofluorescent antibody (FA) test kits for detection of Cryptosporidium oocysts of species other
than Cryptosporidium parvum. Am J Trop Med Hyg 54, 274–279 (1996).
49. Yu, J.R., O’Hara, S.P., Lin, J.L., Dailey, M.E., and Cain, G. A common oocyst surface antigen of
Cryptosporidium recognized by monoclonal antibodies. Parasitol Res 88, 412–420 (2002).
50. Church, D. et al. Screening for Giardia/Cryptosporidium infections using an enzyme immunoassay in a
centralized regional microbiology laboratory. Arch Pathol Lab Med 129, 754–759 (2005).
51. Garcia, L.S. and Shimizu, R.Y. Evaluation of nine immunoassay kits (enzyme immunoassay and direct
fluorescence) for detection of Giardia lamblia and Cryptosporidium parvum in human fecal specimens.
J Clin Microbiol 35, 1526–1529 (1997).
52. Gaafar, M.R. Evaluation of enzyme immunoassay techniques for diagnosis of the most common intestinal protozoa in fecal samples. Int J Infect Dis 15, e541–e544 (2011).
53. Bialek, R. et al. Comparison of fluorescence, antigen and PCR assays to detect Cryptosporidium parvum
in fecal specimens. Diagn Microbiol Infect Dis 43, 283–288 (2002).
54. Srijan, A. et al. Re-evaluation of commercially available enzyme-linked immunosorbent assay for the
detection of Giardia lamblia and Cryptosporidium spp from stool specimens. Southeast Asian J Trop
Med Public Health 36(Suppl. 4), 26–29 (2005).
55. Chalmers, R.M., Campbell, B.M., Crouch, N., Charlett, A., and Davies, A.P. Comparison of diagnostic sensitivity and specificity of seven Cryptosporidium assays used in the UK. J Med Microbiol
60, 1598–1604 (2011).
56. Katanik, M.T., Schneider, S.K., Rosenblatt, J.E., Hall, G.S., and Procop, G.W. Evaluation of
ColorPAC Giardia/Cryptosporidium rapid assay and ProSpecT Giardia/Cryptosporidium microplate assay for detection of Giardia and Cryptosporidium in fecal specimens. J Clin Microbiol
39, 4523–4525 (2001).
Biology of Foodborne Parasites
57. Garcia, L.S. and Shimizu, R.Y. Detection of Giardia lamblia and Cryptosporidium parvum antigens in
human fecal specimens using the ColorPAC combination rapid solid-phase qualitative immunochromatographic assay. J Clin Microbiol 38, 1267–1268 (2000).
58. Garcia, L.S., Shimizu, R.Y., Novak, S., Carroll, M., and Chan, F. Commercial assay for detection of
Giardia lamblia and Cryptosporidium parvum antigens in human fecal specimens by rapid solid-phase
qualitative immunochromatography. J Clin Microbiol 41, 209–212 (2003).
59. Abdel Hameed, D.M., Elwakil, H.S., and Ahmed, M.A. A single-step immunochromatographic lateralflow assay for detection of Giardia lamblia and Cryptosporidium parvum antigens in human fecal samples. J Egypt Soc Parasitol 38, 797–804 (2008).
60. Regnath, T., Klemm, T., and Ignatius, R. Rapid and accurate detection of Giardia lamblia and
Cryptosporidium spp. antigens in human fecal specimens by new commercially available qualitative
immunochromatographic assays. Eur J Clin Microbiol Infect Dis 25, 807–809 (2006).
61. El-Moamly, A.A. and El-Sweify, M.A. ImmunoCard STAT! cartridge antigen detection assay compared
to microplate enzyme immunoassay and modified Kinyoun’s acid-fast staining technique for detection of
Cryptosporidium in fecal specimens. Parasitol Res 110, 1037–1041 (2012).
62. Weitzel, T., Dittrich, S., Mohl, I., Adusu, E., and Jelinek, T. Evaluation of seven commercial antigen
detection tests for Giardia and Cryptosporidium in stool samples. Clin Microbiol Infect 12, 656–659
63. Goni, P. et al. Evaluation of an immunochromatographic dip strip test for simultaneous detection of
Cryptosporidium spp., Giardia duodenalis, and Entamoeba histolytica antigens in human faecal samples.
Eur J Clin Microbiol Infect Dis 31, 2077–2082 (2012).
64. Agnamey, P. et al. Evaluation of four commercial rapid immunochromatographic assays for detection of
Cryptosporidium antigens in stool samples: A blind multicenter trial. J Clin Microbiol 49, 1605–1607
65. Robinson, T.J., Cebelinski, E.A., Taylor, C., and Smith, K.E. Evaluation of the positive predictive value
of rapid assays used by clinical laboratories in Minnesota for the diagnosis of cryptosporidiosis. Clin
Infect Dis 50, e53–e55 (2010).
66. Xiao, L. et al. Phylogenetic analysis of Cryptosporidium parasites based on the small-subunit rRNA gene
locus. Appl Environ Microbiol 65, 1578–1583 (1999).
67. Sturbaum, G.D. et al. Species-specific, nested PCR-restriction fragment length polymorphism detection
of single Cryptosporidium parvum oocysts. Appl Environ Microbiol 67, 2665–2668 (2001).
68. Amar, C.F., Dear, P.H., and McLauchlin, J. Detection and identification by real time PCR/RFLP analyses
of Cryptosporidium species from human faeces. Lett Appl Microbiol 38, 217–222 (2004).
69. Nichols, R.A., Campbell, B.M., and Smith, H.V. Identification of Cryptosporidium spp. oocysts in United
Kingdom noncarbonated natural mineral waters and drinking waters by using a modified nested PCRrestriction fragment length polymorphism assay. Appl Environ Microbiol 69, 4183–4189 (2003).
70. Coupe, S., Sarfati, C., Hamane, S., and Derouin, F. Detection of Cryptosporidium and identification
to the species level by nested PCR and restriction fragment length polymorphism. J Clin Microbiol
43, 1017–1023 (2005).
71. Jiang, J. and Xiao, L. An evaluation of molecular diagnostic tools for the detection and differentiation of
human-pathogenic Cryptosporidium spp. J Eukaryot Microbiol 50(Suppl.), 542–547 (2003).
72. Alves, M. et al. Subgenotype analysis of Cryptosporidium isolates from humans, cattle, and zoo ruminants in Portugal. J Clin Microbiol 41, 2744–2747 (2003).
73. Sulaiman, I.M. et al. Unique endemicity of cryptosporidiosis in children in Kuwait. J Clin Microbiol
43, 2805–2809 (2005).
74. Widmer, G. Meta-analysis of a polymorphic surface glycoprotein of the parasitic protozoa Cryptosporidium
parvum and Cryptosporidium hominis. Epidemiol Infect 137, 1800–1808 (2009).
75. Feng, Y., Lal, A.A., Li, N., and Xiao, L. Subtypes of Cryptosporidium spp. in mice and other small mammals. Exp Parasitol 127, 238–242 (2011).
76. Gatei, W. et al. Development of a multilocus sequence typing tool for Cryptosporidium hominis.
J Eukaryot Microbiol 53, S43–S48 (2006).
77. Tanriverdi, S. et al. Emergence of distinct genotypes of Cryptosporidium parvum in structured host populations. Appl Environ Microbiol 72, 2507–2513 (2006).
78. Glaberman, S. et al. Three drinking-water-associated cryptosporidiosis outbreaks, Northern Ireland.
Emerg Infect Dis 8, 631–633 (2002).
79. Leoni, F., Gallimore, C.I., Green, J., and McLauchlin, J. Molecular epidemiological analysis of
Cryptosporidium isolates from humans and animals by using a heteroduplex mobility assay and nucleic
acid sequencing based on a small double-stranded RNA element. J Clin Microbiol 41, 981–992 (2003).
80. Chalmers, R.M. et al. Direct comparison of selected methods for genetic categorisation of Cryptosporidium
parvum and Cryptosporidium hominis species. Int J Parasitol 35, 397–410 (2005).
81. Xiao, L. and Ryan, U.M. Molecular epidemiology. In Cryptosporidium and Cryptosporidiosis (Fayer, R.
and Xiao, L., eds.), pp. 119–171 (CRC Press and IWA Publishing, Boca Raton, FL, 2008).
82. Blackburn, B.G. et al. Cryptosporidiosis associated with ozonated apple cider. Emerg Infect Dis
12, 684–686 (2006).
83. Waldron, L.S. et al. Molecular epidemiology and spatial distribution of a waterborne cryptosporidiosis
outbreak in Australia. Appl Environ Microbiol 77, 7766–7771 (2011).
84. Mayne, D.J. et al. A community outbreak of cryptosporidiosis in sydney associated with a public swimming facility: A case-control study. Interdiscip Perspect Infect Dis 2011, 341065 (2011).
85. Ng, J.S., Pingault, N., Gibbs, R., Koehler, A., and Ryan, U. Molecular characterisation of Cryptosporidium
outbreaks in Western and South Australia. Exp Parasitol 125, 325–328 (2010).
86. Chalmers, R.M. et al. Detection of Cryptosporidium species and sources of contamination with
Cryptosporidium hominis during a waterborne outbreak in north west Wales. J Water Health 8, 311–325
87. Valderrama, A.L. et al. Multiple risk factors associated with a large statewide increase in cryptosporidiosis. Epidemiol Infect 137, 1781–1788 (2009).
88. Feng, Y. et al. Extended outbreak of cryptosporidiosis in a pediatric hospital, China. Emerg Infect Dis
18, 312–314 (2012).
89. Insulander, M. et al. Molecular epidemiology and clinical manifestations of human cryptosporidiosis in
Sweden. Epidemiol Infect 141, 1009–1020 (2013).
90. Gherasim, A. et al. Two geographically separated food-borne outbreaks in Sweden linked by an unusual
Cryptosporidium parvum subtype, October 2010. Euro Surveill 17, pii: 20318 (2012).
91. Centers for Disease Control and Prevention. Cryptosporidiosis outbreak at a summer camp—North
Carolina, 2009. MMWR 60, 918–922 (2011).
92. Fournet, N. et al. Simultaneous increase of Cryptosporidium infections in the Netherlands, the United
Kingdom and Germany in late summer season, 2012. Euro Surveill 18, pii:20348 (2013).
93. Elwin, K., Hadfield, S.J., Robinson, G., Crouch, N.D., and Chalmers, R.M. Cryptosporidium viatorum
n. sp. (Apicomplexa: Cryptosporidiidae) among travellers returning to Great Britain from the Indian
subcontinent, 2007–2011. Int J Parasitol 42, 675–682 (2012).
94. Elwin, K., Hadfield, S.J., Robinson, G., and Chalmers, R.M. The epidemiology of sporadic human infections with unusual cryptosporidia detected during routine typing in England and Wales, 2000–2008.
Epidemiol Infect 140, 673–683 (2012).
95. Nichols, G.L. et al. Cryptosporidiosis: A report on the surveillance and epidemiology of Cryptosporidium
infection in England and Wales. Drinking Water Directorate Contract Number DWI 70/2/201, Vol. 142.
Drinking Water Inspectorate, U.K. (2006).
96. Raskova, V. et al. Human cryptosporidiosis caused by Cryptosporidium tyzzeri and C. parvum isolates
presumably transmitted from wild mice. J Clin Microbiol 51, 360–362 (2013).
97. Kvac, M. et al. Gastroenteritis caused by the Cryptosporidium hedgehog genotype in an immunocompetent man. J Clin Microbiol 52, 347–349 (2014).
98. Mor, S.M., DeMaria, A., Jr., Griffiths, J.K., and Naumova, E.N. Cryptosporidiosis in the elderly population of the United States. Clin Infect Dis 48, 698–705 (2009).
99. Dietz, V. et al. Active, multisite, laboratory-based surveillance for Cryptosporidium parvum. Am J Trop
Med Hyg 62, 368–372 (2000).
100. Roy, S.L. et al. Risk factors for sporadic cryptosporidiosis among immunocompetent persons in the
United States from 1999 to 2001. J Clin Microbiol 42, 2944–2951 (2004).
101. Goh, S. et al. Sporadic cryptosporidiosis, North Cumbria, England, 1996–2000. Emerg Infect Dis
10, 1007–1015 (2004).
102. McLauchlin, J. et al. Polymerase chain reaction-based diagnosis of infection with Cryptosporidium in
children with primary immunodeficiencies. Pediatr Infect Dis J 22, 329–335 (2003).
103. Hunter, P.R. and Nichols, G. Epidemiology and clinical features of Cryptosporidium infection in immunocompromised patients. Clin Microbiol Rev 15, 145–154 (2002).
Biology of Foodborne Parasites
104. Wolska-Kusnierz, B. et al. Cryptosporidium infection in patients with primary immunodeficiencies.
J Pediatr Gastroenterol Nutr 45, 458–464 (2007).
105. Krause, I. et al. Cryptosporidiosis in children following solid organ transplantation. Pediatr Infect Dis
J 31, 1135–1138 (2012).
106. Tran, M.Q. et al. Cryptosporidium infection in renal transplant patients. Clin Nephrol 63, 305–309 (2005).
107. Seyrafian, S., Pestehchian, N., Kerdegari, M., Yousefi, H.A., and Bastani, B. Prevalence rate of
Cryptosporidium infection in hemodialysis patients in Iran. Hemodial Int 10, 375–379 (2006).
108. Bonatti, H., Barroso, L.F., 2nd, Sawyer, R.G., Kotton, C.N., and Sifri, C.D. Cryptosporidium enteritis in
solid organ transplant recipients: Multicenter retrospective evaluation of 10 cases reveals an association
with elevated tacrolimus concentrations. Transpl Infect Dis 14, 635–648 (2012).
109. Bandin, F. et al. Cryptosporidiosis in paediatric renal transplantation. Pediatr Nephrol 24, 2245–2255
110. Hunter, P.R. et al. Sporadic cryptosporidiosis case-control study with genotyping. Emerg Infect Dis
10, 1241–1249 (2004).
111. Pollock, K.G. et al. Spatial and temporal epidemiology of sporadic human cryptosporidiosis in Scotland.
Zoonoses Public Health 57, 487–492 (2010).
112. Pintar, K.D. et al. A modified case-control study of cryptosporidiosis (using non-Cryptosporidiuminfected enteric cases as controls) in a community setting. Epidemiol Infect 137, 1789–1799 (2009).
113. Lake, I.R. et al. Case-control study of environmental and social factors influencing cryptosporidiosis. Eur
J Epidemiol 22, 805–811 (2007).
114. Snel, S.J., Baker, M.G., and Venugopal, K. The epidemiology of cryptosporidiosis in New Zealand,
1997–2006. NZ Med J 122, 47–61 (2009).
115. Learmonth, J.J., Ionas, G., Ebbett, K.A., and Kwan, E.S. Genetic characterization and transmission
cycles of Cryptosporidium species isolated from humans in New Zealand. Appl Environ Microbiol
70, 3973–3978 (2004).
116. Chalmers, R.M., Elwin, K., Thomas, A.L., Guy, E.C., and Mason, B. Long-term Cryptosporidium typing reveals the aetiology and species-specific epidemiology of human cryptosporidiosis in England and
Wales, 2000 to 2003. Euro Surveill 14, pii: 19086 (2009).
117. Nazemalhosseini-Mojarad, E., Feng, Y., and Xiao, L. The importance of subtype analysis of
Cryptosporidium spp. in epidemiological investigations of human cryptosporidiosis in Iran and other
Mideast countries. Gastroenterol Hepatol Bed Bench 5, 67–70 (2012).
118. Hunter, P.R. et al. Subtypes of Cryptosporidium parvum in humans and disease risk. Emerg Infect Dis
13, 82–88 (2007).
119. Mallon, M.E., MacLeod, A., Wastling, J.M., Smith, H., and Tait, A. Multilocus genotyping of
Cryptosporidium parvum type 2: Population genetics and sub-structuring. Infect Genet Evol 3, 207–218
120. Baldursson, S. and Karanis, P. Waterborne transmission of protozoan parasites: Review of worldwide
outbreaks—An update 2004–2010. Water Res 45, 6603–6614 (2011).
121. McLauchlin, J., Amar, C., Pedraza-Diaz, S., and Nichols, G.L. Molecular epidemiological analysis of Cryptosporidium spp. in the United Kingdom: Results of genotyping Cryptosporidium spp. in
1,705 fecal samples from humans and 105 fecal samples from livestock animals. J Clin Microbiol
38, 3984–3990 (2000).
122. Hlavsa, M.C. et al. Surveillance for waterborne disease outbreaks and other health events associated with
recreational water—United States, 2007–2008. MMWR Surveill Summ 60, 1–32 (2011).
123. Gallaher, M.M. et al. Cryptosporidiosis and surface water. Am J Public Health 79, 39–42 (1989).
124. Frost, F.J. et al. Serological responses to Cryptosporidium antigens among users of surface- vs. groundwater sources. Epidemiol Infect 131, 1131–1138 (2003).
125. Khalakdina, A., Vugia, D.J., Nadle, J., Rothrock, G.A., and Colford, J.M., Jr. Is drinking water a risk factor for endemic cryptosporidiosis? A case-control study in the immunocompetent general population of
the San Francisco Bay Area. BMC Public Health 3, 11 (2003).
126. Sorvillo, F. et al. Municipal drinking water and cryptosporidiosis among persons with AIDS in Los
Angeles County. Epidemiol Infect 113, 313–320 (1994).
127. Pintar, K.D. et al. Considering the risk of Infection by Cryptosporidium via consumption of municipally
treated drinking water from a surface water source in a Southwestern Ontario community. Risk Anal
32, 1122–1138 (2012).
128. Fayer, R. Cryptosporidium: A water-borne zoonotic parasite. Vet Parasitol 126, 37–56 (2004).
129. Karanis, P., Kourenti, C., and Smith, H. Waterborne transmission of protozoan parasites: A worldwide
review of outbreaks and lessons learnt. J Water Health 5, 1–38 (2007).
130. Semenza, J.C. and Nichols, G. Cryptosporidiosis surveillance and water-borne outbreaks in Europe. Euro
Surveill 12, E13–E14 (2007).
131. USEPA. National primary drinking water regulations: Long term 2 enhanced surface water treatment
rule. Federal Register 71, 654–786 (2006).
132. May, A. The benefits of drinking water quality regulation—England and Wales. Water Sci Technol
54, 387–393 (2006).
133. Gostin, L.O., Lazzarini, Z., Neslund, V.S., and Osterholm, M.T. Water quality laws and waterborne diseases: Cryptosporidium and other emerging pathogens. Am J Public Health 90, 847–853 (2000).
134. Chalmers, R.M. et al. Cryptosporidium sp. rabbit genotype, a newly identified human pathogen. Emerg
Infect Dis 15, 829–830 (2009).
135. Tumwine, J.K. et al. Cryptosporidium parvum in children with diarrhea in Mulago Hospital, Kampala,
Uganda. Am J Trop Med Hyg 68, 710–715 (2003).
136. Peng, M.M. et al. Molecular epidemiology of cryptosporidiosis in children in Malawi. J Eukaryot
Microbiol 50(Suppl.), 557–559 (2003).
137. Bhattacharya, M.K., Teka, T., Faruque, A.S., and Fuchs, G.J. Cryptosporidium infection in children in
urban Bangladesh. J Trop Pediatr 43, 282–286 (1997).
138. Leach, C.T., Koo, F.C., Kuhls, T.L., Hilsenbeck, S.G., and Jenson, H.B. Prevalence of Cryptosporidium
parvum infection in children along the Texas-Mexico border and associated risk factors. Am J Trop Med
Hyg 62, 656–661 (2000).
139. Robertson, L.J. and Gjerde, B. Occurrence of parasites on fruits and vegetables in Norway. J Food Prot
64, 1793–1798 (2001).
140. Fayer, R., Dubey, J.P., and Lindsay, D.S. Zoonotic protozoa: From land to sea. Trends Parasitol
20, 531–536 (2004).
141. Budu-Amoako, E., Greenwood, S.J., Dixon, B.R., Barkema, H.W., and McClure, J.T. Foodborne illness
associated with Cryptosporidium and Giardia from livestock. J Food Prot 74, 1944–1955 (2011).
142. Millard, P.S. et al. An outbreak of cryptosporidiosis from fresh-pressed apple cider [published erratum
appears in JAMA March 8, 1995;273(10):776]. JAMA 272, 1592–1596 (1994).
143. Quiroz, E.S. et al. An outbreak of cryptosporidiosis linked to a foodhandler. J Infect Dis 181, 695–700
144. Ponka, A. et al. A foodborne outbreak due to Cryptosporidium parvum in Helsinki, November 2008.
Euro Surveill 14, pii: 19269 (2009).
145. Ethelberg, S. et al. A foodborne outbreak of Cryptosporidium hominis infection. Epidemiol Infect
137, 348–356 (2009).
146. Yoshida, H. et al. An outbreak of cryptosporidiosis suspected to be related to contaminated food, October
2006, Sakai City, Japan. Jpn J Infect Dis 60, 405–407 (2007).
147. Pereira, M.D., Atwill, E.R., Barbosa, A.P., Silva, S.A., and Garcia-Zapata, M.T. Intra-familial and extrafamilial risk factors associated with Cryptosporidium parvum infection among children hospitalized for
diarrhea in Goiania, Goias, Brazil. Am J Trop Med Hyg 66, 787–793 (2002).
148. Robertson, B. et al. Case-control studies of sporadic cryptosporidiosis in Melbourne and Adelaide,
Australia. Epidemiol Infect 128, 419–431 (2002).
149. Bern, C. et al. Epidemiologic differences between cyclosporiasis and cryptosporidiosis in Peruvian children. Emerg Infect Dis 8, 581–585 (2002).
150. Cama, V.A. et al. Cryptosporidium species and subtypes and clinical manifestations in children, Peru.
Emerg Infect Dis 14, 1567–1574 (2008).
151. Creek, T.L. et al. Hospitalization and mortality among primarily nonbreastfed children during a large outbreak of diarrhea and malnutrition in Botswana, 2006. J Acquir Immune Defic Syndr 53, 14–19 (2010).
152. Hunter, P.R. et al. Health sequelae of human cryptosporidiosis in immunocompetent patients. Clin Infect
Dis 39, 504–510 (2004).
153. De Angelis, C. et al. An update on AIDS-related cholangiopathy. Minerva Gastroenterol Dietol
55, 79–82 (2009).
154. Manabe, Y.C. et al. Cryptosporidiosis in patients with AIDS: Correlates of disease and survival. Clin
Infect Dis 27, 536–542 (1998).
Biology of Foodborne Parasites
155. Clayton, F., Heller, T., and Kotler, D.P. Variation in the enteric distribution of cryptosporidia in acquired
immunodeficiency syndrome. Am J Clin Pathol 102, 420–425 (1994).
156. Lumadue, J.A. et al. A clinicopathologic analysis of AIDS-related cryptosporidiosis. AIDS 12, 2459–
2466 (1998).
157. Cama, V.A. et al. Differences in clinical manifestations among Cryptosporidium species and subtypes in
HIV-infected persons. J Infect Dis 196, 684–691 (2007).
158. Pantenburg, B. et al. Intestinal immune response to human Cryptosporidium sp. infection. Infect Immun
76, 23–29 (2008).
159. Hernandez, J. et al. Substance P is responsible for physiological alterations such as increased chloride ion
secretion and glucose malabsorption in cryptosporidiosis. Infect Immun 75, 1137–1143 (2007).
160. Pantenburg, B. et al. Human CD8(+) T cells clear Cryptosporidium parvum from infected intestinal epithelial cells. Am J Trop Med Hyg 82, 600–607 (2010).
161. Costa, L.B. et al. Cryptosporidium–malnutrition interactions: Mucosal disruption, cytokines and TLR
signaling in a weaned murine model. J Parasitol 97, 1113–1120 (2011).
162. Zhou, R., Gong, A.Y., Eischeid, A.N., and Chen, X.M. miR-27b targets KSRP to coordinate TLR4mediated epithelial defense against Cryptosporidium parvum infection. PLoS Pathog 8, e1002702
163. Gargala, G. Drug treatment and novel drug target against Cryptosporidium. Parasite 15, 275–281 (2008).
164. Rossignol, J.F. Cryptosporidium and Giardia: Treatment options and prospects for new drugs. Exp
Parasitol 124, 45–53 (2010).
165. Pantenburg, B., Cabada, M.M., and White, A.C., Jr. Treatment of cryptosporidiosis. Expert Rev Anti
Infect Ther 7, 385–391 (2009).
166. Abubakar, I., Aliyu, S.H., Arumugam, C., Usman, N.K., and Hunter, P.R. Treatment of cryptosporidiosis
in immunocompromised individuals: Systematic review and meta-analysis. Br J Clin Pharmacol 63,
387–393 (2007).
167. Bailey, J.M. and Erramouspe, J. Nitazoxanide treatment for giardiasis and cryptosporidiosis in children.
Ann Pharmacother 38, 634–640 (2004).
168. Zardi, E.M., Picardi, A., and Afeltra, A. Treatment of cryptosporidiosis in immunocompromised hosts.
Chemotherapy 51, 193–196 (2005).
169. Werneck-Silva, A.L. and Prado, I.B. Gastroduodenal opportunistic infections and dyspepsia in HIVinfected patients in the era of Highly Active Antiretroviral Therapy. J Gastroenterol Hepatol 24, 135–139
170. Maggi, P. et al. Effect of antiretroviral therapy on cryptosporidiosis and microsporidiosis in patients
infected with human immunodeficiency virus type 1. Eur J Clin Microbiol Infect Dis 19, 213–217 (2000).
171. Certad, G. et al. Cryptosporidiosis in HIV-infected Venezuelan adults is strongly associated with acute or
chronic diarrhea. Am J Trop Med Hyg 73, 54–57 (2005).
172. Tuli, L., Gulati, A.K., Sundar, S., and Mohapatra, T.M. Correlation between CD4 counts of HIV patients
and enteric protozoan in different seasons—An experience of a tertiary care hospital in Varanasi (India).
BMC Gastroenterol 8, 36 (2008).
173. Juranek, D.D. Cryptosporidiosis: Sources of infection and guidelines for prevention. Clin Infect Dis
21, S57–S61 (1995).
174. Kaplan, J.E. et al. Guidelines for prevention and treatment of opportunistic infections in HIV-infected
adults and adolescents: Recommendations from CDC, the National Institutes of Health, and the HIV
Medicine Association of the Infectious Diseases Society of America. MMWR Recomm Rep 58, 1–207;
quiz CE1–CE4 (2009).
175. Palmer, C.J. et al. Cryptosporidium muris, a rodent pathogen, recovered from a human in Peru. Emerg
Infect Dis 9, 1174–1176 (2003).
176. Upton, S.J. and Current, W.L. The species of Cryptosporidium (Apicomplexa: Cryptosporidiidae) infecting mammals. J Parasitol 71, 625–629 (1985).
177. Tilley, M., Upton, S.J., and Chrisp, C.E. A comparative study on the biology of Cryptosporidium sp. from
guinea pigs and Cryptosporidium parvum (Apicomplexa). Can J Microbiol 37, 949–952 (1991).
178. Sreter, T. et al. Morphologic, host specificity, and molecular characterization of a Hungarian
Cryptosporidium meleagridis isolate. Appl Environ Microbiol 66, 735–738 (2000).
179. Lindsay, D.S. et al. Cryptosporidium andersoni n. sp. (Apicomplexa: Cryptosporiidae) from cattle, Bos
taurus. J Eukaryot Microbiol 47, 91–95 (2000).
180. Fayer, R. et al. Cryptosporidium canis n. sp. from domestic dogs. J Parasitol 87, 1415–1422 (2001).
181. Morgan-Ryan, U.M. et al. Cryptosporidium hominis n. sp. (Apicomplexa: Cryptosporidiidae) from
Homo sapiens. J Eukaryot Microbiol 49, 433–440 (2002).
182. Ryan, U.M. et al. Cryptosporidium suis n. sp. (Apicomplexa: Cryptosporidiidae) in pigs (Sus scrofa).
J Parasitol 90, 769–773 (2004).
183. Fayer, R., Santin, M., and Xiao, L. Cryptosporidium bovis n. sp. (Apicomplexa: Cryptosporidiidae) in
cattle (Bos taurus). J Parasitol 91, 624–629 (2005).
184. Ryan, U.M., Power, M., and Xiao, L. Cryptosporidium fayeri n. sp. (Apicomplexa: Cryptosporidiidae)
from the Red Kangaroo (Macropus rufus). J Eukaryot Microbiol 55, 22–26 (2008).
185. Power, M.L. and Ryan, U.M. A new species of Cryptosporidium (Apicomplexa: Cryptosporidiidae) from
Eastern Grey Kangaroos (Macropus giganteus). J Parasitol 94, 1114–1117 (2008).
186. Fayer, R., Santin, M., and Trout, J.M. Cryptosporidium ryanae n. sp (Apicomplexa: Cryptosporidiidae)
in cattle (Bos taurus). Vet Parasitol 156, 191–198 (2008).
187. Fayer, R. and Santin, M. Cryptosporidium xiaoi n. sp. (Apicomplexa: Cryptosporidiidae) in sheep
(Ovis aries). Vet Parasitol 146, 192–200 (2009).
188. Fayer, R., Santin, M., and Macarisin, D. Cryptosporidium ubiquitum n. sp. in animals and humans.
Vet Parasitol 172, 23–32 (2010).
189. Robinson, G. et al. Re-description of Cryptosporidium cuniculus (Apicomplexa: Cryptosporidiidae):
Morphology, biology and phylogeny. Int J Parasitol 40, 1539–1548 (2010).
190. Ren, X. et al. Cryptosporidium tyzzeri n. sp. (Apicomplexa: Cryptosporidiidae) in domestic mice (Mus
musculus). Exp Parasitol 130, 274–281 (2012).
191. Kvac, M. et al. Cryptosporidium scrofarum n. sp. (Apicomplexa: Cryptosporidiidae) in domestic pigs
(Sus scrofa). Vet Parasitol 191, 218–227 (2013).
192. Current, W.L., Upton, S.J., and Haynes, T.B. The life cycle of Cryptosporidium baileyi n. sp.
(Apicomplexa: Cryptosporidiidae) infecting chickens. J Protozool 33, 289–296 (1986).
193. Ryan, U.M. et al. A redescription of Cryptosporidium galli Pavlasek, 1999 (Apicomplexa:
Cryptosporidiidae) from birds. J Parasitol 89, 809–813 (2003).
194. Jirku, M. et al. New species of Cryptosporidium Tyzzer, 1907 (Apicomplexa) from amphibian host:
Morphology, biology and phylogeny. Folia Parasitol (Praha) 55, 81–94 (2008).
195. Xiao, L. et al. Genetic diversity of Cryptosporidium spp. in captive reptiles. Appl Environ Microbiol
70, 891–899 (2004).
196. Alvarez-Pellitero, P. and Sitja-Bobadilla, A. Cryptosporidium molnari n. sp. (Apicomplexa:
Cryptosporidiidae) infecting two marine fish species, Sparus aurata L. and Dicentrarchus labrax L. Int
J Parasitol 32, 1007–1021 (2002).
197. Zhu, G. and Xiao, L. Cryptosporidium species. In Genomes of Foodborne and Waterborne Pathogens
(Fratamico, P., Liu, Y., and Kathariou, S., eds.), pp. 271–286 (American Society for Microbiology,
Washington, DC, 2011).
Cyclospora cayetanensis
Ynes R. Ortega and Jeevan B. Sherchand
6.1 Introduction..................................................................................................................................... 97
6.2 Morphology and Classification....................................................................................................... 97
6.3 Biology, Genetics, and Genomics................................................................................................. 100
6.4 Diagnosis and Typing.................................................................................................................... 100
6.5 Epidemiology and Molecular Epidemiology.................................................................................101
6.6 Pathogenesis and Clinical Features.............................................................................................. 104
6.7 Treatment and Prevention............................................................................................................. 106
6.8 Conclusions................................................................................................................................... 107
References............................................................................................................................................... 107
6.1 Introduction
In the late 1980s and early 1990s, a few reports described cases of gastroenteritis caused by an unknown
organism in travelers returning from developing countries. It was also identified as a cause of diarrhea in children and adults of these countries. These structures were called coccidian-like bodies,
cyanobacteria-like organisms, etc. Their characteristics were similar to Cryptosporidium, another
Apicomplexan parasite that also affects children and travelers. In 1992, Ortega et al. characterized and
named this organism as Cyclospora cayetanensis. Although some reports described Cyclospora as a
causative agent of travelers’ diarrhea, in 1995 the first U.S. outbreak of Cyclospora occurred in nontravelers. Since then, several Cyclospora outbreaks were linked to imported berries and vegetables that were
consumed raw. This chapter describes the biology of Cyclospora, its epidemiology, detection, pathogenesis, prevention, and their major implications in food trade.
6.2 Morphology and Classification
As all coccidian parasites, their life cycles are complex and have two cycles of multiplication with various stages. The asexual and sexual stages of C. cayetanensis were described in morphologic and electron
microscopic studies.1 Overall, Cyclospora has polymorphic intracellular stages in its life cycle.
The infective stage of this coccidian is the sporulated oocyst that is the environmentally resistant
cystic stage. Only unsporulated oocysts are excreted in the feces of the infected individual. Unlike some
other coccidia that infect humans, Cyclospora oocysts require days outside the host after being passed in
bowel movements to develop into an infectious stage. The immature oocyst is spheroidal with a diameter
of 8–10 μm (7.7–9.9 μm) (Figure 6.1a). This oocyst has a 113 nm thick bilayered wall. The outer rough
coat is 63 nm thick, while the inner smooth layer is 50 nm thick. A polar body and oocyst residuum
are present. The unsporulated oocysts may have a granular cytoplasm. The oocysts appear as nonrefractile, round, hyaline structures containing an arrangement of refractile membrane with six to nine
Biology of Foodborne Parasites
FIGURE 6.1 C. cayetanensis oocysts. (a) Autofluorescence. (b) Unsporulated oocyst. (c) Sporulated oocyst. (d) Safraninstained oocysts. (e) Modified acid-fast–stained oocyst.
bound globules. The mature oocysts (Figure 6.1c) have a fibrillar coat and cell wall similar to that of the
unsporulated oocysts.2 A diagram of the life cycle of Cyclospora is presented in Figure 6.2.
Each sporulated oocyst contains two ovoidal sporocysts measuring 6.3 μm long and 4.0 μm wide. Each
sporocyst has a 62 nm thick wall. The sporocyst has Stieda and sub-Stieda bodies. Sporocyst residuum
has large spherical globules.2 Each sporocyst has two sporozoites with a total of four sporozoites per
oocyst. The size of each sporozoite is 9.0 μm long and 1.2 μm wide.2 They are slender and crescent- or
spindle-shaped, are motile, and are infectious. The trophozoite is intracellular and measures about 4 μm
long and 2 μm wide in size.1
The asexual multiplication occurs during the meront phase. There are two types of meronts: the first
meront (meront I) having 8–12 merozoites and the second meront (meront II) having 4 merozoites. These
meronts are about 3–4 μm long and 0.5 μm wide.3 The merozoites are crescent shaped.4
The sexual multiplication occurs when the meront II forms gamonts known as microgametocytes or macrogametocytes. The microgametocytes, which are the male gamonts, are filled with numerous flagellated
sperm-like motile microgametes. Most of the meront II form female gamonts known as macrogamonts or
macrogametocytes. They are larger in size than microgamonts. As result of the fertilization of the macrogametocyte by the microgametocyte, the zygote is formed and differentiates to form the immature oocyst.
Cyclospora is classified based on its morphological characteristics of the various stages of the life cycle.
Toxoplasma and Cystoisospora also belong to the Coccidiasina subclass but belong to different families. In recent years, several Cyclospora spp. have been identified in other animal species in addition to
humans (Table 6.1).
Domain: Eukaryota
Kingdom: Chromalveolata
Superphylum: Alveolata
Cyclospora cayetanensis
5 Ingestion of
Sporulated oocysts
enter the food chain
Oocyst sporulation
in the environment
2 Environmental
i = Infective stage
d = Diagnostic stage
1 d
Excretion of
oocysts in
the stool
Meront Meront
FIGURE 6.2 Life cycle of Cyclospora. (From
Cyclospora Species Recently Described Morphologically Similar to C. cayetanensis
Cyclospora Species
C. cayetanensis
C. cercopitheci
C. colobi
C. papionis
Cyclospora sp.
Host Common Name
Homo sapiens
Cercopithecus aethiops
Colobus guereza
Papio anubis
Rhinopithecus roxellana
African green monkeys
Colobus monkeys
Golden snub-nosed monkeys
Ortega et al. [2]
Eberhard et al. [5a]
Eberhard et al. [5a]
Eberhard et al. [5a]
Zhao et al. [5]
Biology of Foodborne Parasites
Phylum: Apicomplexa
Class: Conoidasida
Subclass: Coccidiasina
Order: Eucoccidiorida
Suborder: Eimeriorina
Family: Eimeriidae
Genus: Cyclospora
Species: Cyclospora cayetanensis
6.3 Biology, Genetics, and Genomics
The life cycle of Cyclospora seems to be similar to that of other coccidia. As of now, it seems to require
one host where it will infect, propagate asexually and sexually, resulting in a formation of the environmentally resistant oocyst.
The marked seasonality of Cyclospora is very specific in each endemic region. The environmental conditions that are responsible for this seasonality are unknown; neither are reasons for the oocyst
­survival for months (considered the low season). No reservoirs or intermediary hosts have been identified yet. In a recent study in China, seasonality was also observed in snub-nosed monkeys infected
with Cyclospora sp.5 Unlike C. cayetanensis, the other three Cyclospora species of nonhuman primates
(C. colobi and C. cercopitheci) do not show any seasonality.
C. cayetanensis seems to be exclusively anthroponotic. Few reports identified Cyclospora oocysts in
the feces of dogs, chickens, and ducks.6–8 However, many attempts to reproduce the infection in laboratory animals have been unsuccessful.9 Neither intracellular stages in tissues of animals confirming
infection nor amplification of number of oocyst expected after asexual and sexual multiplication of the
parasite has been reported. Since these animal species are coprophagic, identifying oocysts may just
reflect the passage of oocysts through their gastrointestinal tract.
Phylogenetic studies based on the 18S rRNA gene sequences place Cyclospora in the Eimeriidae family within the mammalian and avian Eimeria species. Morphologically though, the Eimeria spp. have
four sporocysts and two sporozoites per sporocyst, whereas Cyclospora has two sporocysts and two
sporozoites per sporocyst.
6.4 Diagnosis and Typing
Diagnosis of cyclosporiasis is done based initially on the clinical presentation (diarrhea, nausea, anorexia,
etc.). Travel history of individuals returning from endemic regions with gastrointestinal illness is also
Fecal samples are examined for the presence of oocysts. Oocysts autofluoresce (Figure 6.1a) when
observed under an epifluorescence microscope10 and the oocyst morphology can be easily i­dentified
by bright-field microscopy in samples with a large number of oocysts (Figure 6.1b). Oocysts can be
concentrated from fecal samples using the ethyl acetate method, discontinuous sucrose gradients, or
cesium chloride. The modified acid-fast stains such as the modified safranin technique with m
­ icrowave
heating11 (Figure 6.1d) and the modified Ziehl–Neelsen staining12 (Figure 6.1e) can help identify
Cyclospora as well as other coccidian oocysts. These methods are comparable to the autofluorescence observation, but the latter is faster,13 and oocyst identification can be confirmed using phase or
differential interference contrast (DIC) microscopy.11,14,15 Although these methods are relatively easy to
implement in a clinical laboratory, when examining environmental samples, much care should be taken
when choosing a diagnostic method. With these samples, unexperienced microscopists may confuse
artifacts, vegetable parts, pollen, spores, and coccidian oocysts noninfectious to humans as Cyclospora.
Microscopists familiarized with Cyclospora will be able to discriminate C. cayetanensis from artifacts
Cyclospora cayetanensis
or other parasitic organisms. When examining environmental samples, in addition to microscopy,
molecular testing will facilitate the accurate identification of Cyclospora oocysts. Sporulation can also
be done for Cyclospora confirmation (Figure 6.1c).
Primers to amplify the 18S rRNA gene have been useful to detect Cyclospora in clinical samples.
It has also been useful for speciation of Cyclospora infecting nonhuman primates. To test environmental
samples, this PCR assay was complemented with RFLP using the Mnl1 restriction endonuclease to differentiate Cyclospora from other Eimeria spp.16 The development of the oligonucleotide-ligation assay17
facilitated the implementation of the assay by the food regulators and inspectors. Currently, the U.S.
Food and Drug Administration (FDA) uses a method combining a PCR followed by a real-time PCR,
thus increasing the sensitivity of the assay. Outbreak investigation studies are more effective when using
molecular tools.18,19 Other methods targeting the 18S have also been described.20 The ITS-1 and hsp70
genes were also used with the goal to discriminate between isolates and that could be used during traceback studies,21,22 although they have only limited typing capacities.
One of the critical steps for parasite identification (including Cyclospora) is the recovery of oocysts
from food matrices including fruits and vegetables. Various washes have been examined providing different recovery efficiencies.23–26 Shields and collaborators proposed the use of 0.1% Alconox, a laboratory detergent that contains dodecylbenzenesulfonate, a surfactant, and tetrasodium pyrophosphate,
resulting in improved recoveries (72.6% ± 6.6%) compared with deionized water (38.4% ± 10.1%).24
Because much is relied on using molecular tools to detect Cyclospora in food samples, commercial
DNA extraction kits have been frequently used. The FastDNA SPIN Kit for soil, QIAamp DNA Mini
Stool Kit, and UNEX method performed similarly and better than other kits tested.27
6.5 Epidemiology and Molecular Epidemiology
Cyclospora has been described in travelers and in communities of various locations worldwide. Since the
1980s, the first cases corresponded to travelers seeking medical assistance at travel clinics. These were
described as a coccidian parasite, cyanobacteria-like bodies, coccidian-like bodies, big Cryptosporidium, etc.
Cyclospora infections can be acquired in various countries. Most of the cases reported were from
travelers returning from developing countries.13 In few locations worldwide, Cyclospora was studied
more in depth resulting in a better understanding of its epidemiology. Although not exclusive, more studies looking for gastrointestinal etiologies have identified Cyclospora in regions of Nepal, Peru, Haiti,
and Guatemala as endemic. Fewer studies in regions of Mexico, Dominican Republic, China, India,
Colombia, Venezuela, Honduras, and Southeast Asia indicate that these countries may also have areas of
endemicity for Cyclospora.13,28
The marked seasonality of Cyclospora varies by location. In Nepal, the peak season, where most of
the cases are observed, is during May–July. Later peak transmission, July–September, is seen in central
China. In Peru, most of the cases are observed during the months of January–May. In Guatemala, the
high season falls on May–July and in Haiti from February to April. The factors that trigger this seasonality are unknown.
In the United States, most of the cases have been reported during the months of May–August. This
may be associated with the importation of certain food commodities coinciding with the high season
for these exporting countries. In U.S. outbreaks, the implicated foods were not always available nor the
source of these food items. As of now, Cyclospora does not seem to be endemic in the United States, and
the cases reported are either from travelers returning from endemic areas or by ingestion of imported
food items (Table 6.2).
The largest outbreaks in the United States occurred in 1995–1997 and 2013. These outbreaks were
mostly associated with consumption of imported berries. The first-year, domestically grown strawberries
were initially implicated; however, further epidemiological studies demonstrated that the source of the
outbreaks was more likely to be from imported raspberries.29 The first reported waterborne outbreak in
the United States was in 1990 where contaminated water was considered to be the source of infection.
Lettuce, mesclun, basil, snow peas, baby greens, and green onions have been implicated in other foodborne outbreaks of Cyclospora (Table 6.2).
United States (20 states and District of
Columbia); Canada (2 provinces)
United States (13 states and District of
Columbia); Canada (1 provinces)
U.S. cruise ship, Florida departure
United States, Virginia
United States, Georgia
Canada, Ontario
Canada, Ontario
United States, Florida
United States, Philadelphia
Colombia, Medellin
United States, California
United States, Tennessee
United States, Connecticut
United States, Florida
United States, Massachusetts
United States, Illinois
Nepal, Pokhara
Puerto Rico
Turkey, Izmir
Country, City
Date of
21 (1 cluster)
17 (1 cluster)
221 (13 clusters)
38 (2 clusters)
Clinical Cases
Specific Vehicle
Source of Vehicle
Guatemala raspberry, U.S. strawberries,
frozen Chile raspberries
Fruit berry
Raspberries, wedding cake Guatemala
Salads and juice
Mixed berries
Mixed berries
Blackberries and raspberries n/d
Fruit plate
Probable fruit salad
Berry garnish—raspberries
from Guatemala
Dessert (berry)
Tap water
United States
River and municipal waters Nepal
Drinking water
Raspberries country club
Guatemala, Chile
Berry dessert, wedding
California strawberries, Florida
blueberries, Guatemala blackberries,
Guatemala/Chile raspberries
Food Vehicle Associated with Cyclospora Food and Waterborne Outbreaks
Botero-Garces et al.57
(Continued )
Flemming et al.51
Rabold et al.33
Biology of Foodborne Parasites
Canada, British Columbia
United States, Florida
Canada, Sarnia
United States, Florida
Germany, Southwest
Mexico, Monterrey
Cruise ship
United States, Pennsylvania
United States, Wisconsin
United States, New York
United States, Florida
Indonesia, Bogor
Peru, Lima
United States, Illinois
Peru, Lima
Turkey, Istanbul, Kocaeli
United States, New York
United States, Atlanta
United States, 25 states
food item
Northern Virginia, Washington
United States, Washington, D.C.
United States, Missouri
Country, City
Date of
Specific Vehicle
70 available
241 cases
Salad mix? Cilantro?
Fresh produce
Snow peas
Green peas
Salad berros
Basil–pesto pasta salad
341 (57 clusters) Basil–pesto pasta salad
62 (2 events)
Chicken pasta and tomato
basil salad
Thai basil
592 (6 clusters) Fresh basil
Basil, pesto
12 (1 cluster)
Mesclun salad
Salads, green leafy herbs
Clinical Cases
Food Vehicle Associated with Cyclospora Food and Waterborne Outbreaks
TABLE 6.2 (Continued )
United States
Fundraiser event
South France, lettuce; south Italy,
mixed lettuce and herbs; Germany,
Wedding and christening
Basil from Mexico and United States
Source of Vehicle
Torres-Slimming et al.19
Mundaca et al.18
Gibbs et al.49
Ayala-Gaytan et al.63
Hoang et al.60
Doller et al.62
Cyclospora cayetanensis
Biology of Foodborne Parasites
In a recent study in Canada, of a total of 544 retail samples, 9 (1.7%) were positive to Cyclospora by
PCR–RFLP and 7 of these were 100% homologous to Cyclospora by sequencing.30 Most of the examined samples were imported from the United States, Canada, and Mexico; however, none of the samples
from Mexico were positive for Cyclospora.
In 2001–2009, Canada reported 27 produce-related outbreaks. Of those, seven were associated with
Cyclospora. In 2001, 17 cases were reported. Eleven of the 12 cases were exposed to Thai basil produced
in the United States.31 Cilantro was implicated in an outbreak in 2003, sickening 11 people and 8 cases in
2004. Mangos or basil was identified as possible sources of infection of 17 cases of Cyclospora in 2004.
In 2005, basil was implicated with 44 illnesses in one outbreak and 200 cases in a second outbreak.
Again in 2006, basil or garlic was linked to 28 cases of cyclosporiasis.31
In the United States and Canada, raspberries were associated with Cyclospora outbreaks. The largest ones occurred in 1996 with 1465 cases and in 1997 with more than 1000 cases. In 2005, 592 cases
were implicated in a Cyclospora outbreak in Florida13 and more than 100 cases in Atlanta in 2012. The
outbreaks of 1995–1997 showed the need for a trace-back system that could allow identification more
expeditiously the implicated food item, distribution, and source of origin. Still, every year a few cases
and outbreaks occur in the United States, and in most instances the food item is not identified.
Closed and small populations such as military units (which share water and food and sanitary facilities) have been also affected by diarrheal illness outbreaks. In 2011, a U.S. military maneuver enhancement brigade was deployed to San Vicente in El Salvador. Four of 67 soldiers with gastrointestinal illness
developed Cyclospora infection. Consumption of food from local vendors prepared without proper
monitoring was attributed to the source of the infections.32 In Nepal, 12 out of 14 British soldiers and
dependents developed diarrheal illness. Of eight fecal samples examined, six had Cyclospora oocysts.
This outbreak implicated drinking water as the source of infection. The chlorinated water (0.3–0.8 ppm)
they consumed was a mixture of river and municipal water. Although the water did not contain coliform bacteria, it contained Cyclospora oocysts.33 Two other reports describe Cyclospora outbreaks in a
Peruvian Naval Recruit Base. In 2004, 127 recruits developed diarrhea. Cyclospora was identified in 24
of 77 cases (with diarrhea) and 3 of 35 controls (no diarrhea). Fresh vegetables were suspected to be the
source of infection, but this was not confirmed.19 In 2005, another outbreak occurred at the same naval
base. Cyclospora was identified in 20 of 35 cases and in 3 of 15 controls.18
Fresh produce has been mostly associated with foodborne outbreaks of Cyclospora, and even if individuals do not visit endemic regions, foods contaminated with Cyclospora oocysts can cause outbreaks
in unsuspected situations. That is the case of two outbreaks that occurred in 2010 implicating two cruises
and one ship. The first outbreak reported cases days after the passengers arrived in Australia, whereas
the second outbreak occurred while the passengers were in the cruise ship. Cruise 1 had reported 26
confirmed Cyclospora cases and 8 suspected cases with a minimum attack rate of 1.7% (34/2047). In
cruise 2, 46 confirmed cases and 186 suspected cases were reported with a minimum attack rate of
11.5% (232/2010). It was suspected that fresh produce potentially contaminated with Cyclospora oocysts
taken on board at a Southeast Asian port (Malaysia or Singapore) could be responsible for these two
In the past few years, Cyclospora outbreaks were identified, but reports of these have been only available
through the press, web news, or databases such as ProMed. In 2013, the United States again experienced
a large Cyclospora outbreak. A total of 631 cases were reported in 25 states and New York City. Eight
percent of ill persons (49 cases) were hospitalized as result of the Cyclospora infection. Texas (n = 270)
and Iowa (n = 140) had the most cases. The onset of illness is estimated to be from mid-June to early July
(, accessed July 30, 2014).
The source of infection was undermined, but multiple food vehicles from Mexico appeared to be involved.
6.6 Pathogenesis and Clinical Features
The clinical manifestation of Cyclospora infections is similar to that of Cryptosporidium and is characterized by watery explosive diarrhea, anorexia, nausea, and abdominal pain. It can be confused
with Cryptosporidium infections. The severity of the illness seems to be correlated to the repeated
Cyclospora cayetanensis
exposures with the parasite. Asymptomatic infections are common in the local populations of the
endemic locations of countries know to have Cyclospora. Children are usually infected early in life,
and subsequent infections are usually of shorter duration and less severe. In adults, it is infrequent
and if infected they are usually asymptomatic. Asymptomatic infections may also occur in others
including those with HIV infection. Incubation period is usually 1–14 days (mean, 7 days). It may be
preceded by a flu-like prodrome. Low-grade fever and malabsorption may occur. The onset of illness
is abrupt in as many as 30% of cases. The illness is characterized by cyclical diarrhea (explosive
at times; up to numerous times with median of six watery stools per day) accompanied by fatigue,
malaise, anorexia, nausea, weight loss, and abdominal cramping. Diarrhea is self-limiting with the
acute symptoms subsiding within a few days. But it recurs (61% of cases) taking a chronic course
with ­waxing–waning pattern. The illness usually lasts 6–7 weeks but has been reported to persist
for several months if left untreated. Cyclospora infection affects both immunocompetent and immunocompromised individuals, the latter potentially more severely (i.e., chronic, relapsing, protracted
symptoms). Chronic course is more common in HIV cases lasting from several months to a year.
Major signs and symptoms include
• Watery diarrhea (may be explosive) with a median of six stools per day, frequency of diarrhea
possibly many times higher, and no diarrhea in some patients
• Anorexia
• Weight loss (0.9–3.6 kg as reported in one study)
• Fatigue, often pronounced
• Abdominal cramping
• Belching
• Abdominal bloating
• Vomiting
• Low-grade fever (25%)
• Flatulence
Vital signs are normal in most cases. Fever is unusual and, when present, low grade. In the presence
of moderate to severe dehydration, compensatory tachycardia, systolic blood pressure <90 mmHg, and
decreased skin turgor may occur, and the patient may appear ill.
Cyclospora infects the small intestine, particularly the distal duodenum and jejunum. It is unclear
whether the pathogenesis of the disease is due to enterocyte dysfunction or whether toxins are produced. Diarrhea is watery with the absence of any leucocytes or blood, indicating that it may be due
to as yet unidentified toxin. Enterocytes are believed to be invaded by sporozoites, causing release of
cytokines from epithelial cells. Cytokines, in turn, activate and recruit phagocytes from the blood.
These phagocytes release factors such as histamine, prostaglandins, and platelet-aggregating factors
that increase intestinal secretion of chloride and water and inhibit absorption. Another mechanism
of enterocyte damage, besides direct damage caused by the parasite, is ­i nflammation; T cells, proteases, and oxidants secreted from mast cells are responsible for this process. Endoscopic examinations in confirmed cases have provided histological evidence of inflammation leading to small
bowel injury. Acute and chronic inflammation, reactive hyperemia with vascular dilatation and
villous capillary congestion, crypt hyperplasia, epithelial disarray, and partial villous atrophy were
noted in histopathological studies.3,35 Oocysts have been recovered from the duodenal aspirates.
Ultimately, marked destruction of enterocytes causes nutrient malabsorption and increased secretion of fluids and electrolytes from the gut, resulting in secretory and osmotic diarrhea. Abnormal
findings on lactulose or mannitol studies or studies of both have demonstrated intestinal barrier disruption. Abnormal findings on d-xylose studies have demonstrated malabsorption involving proximal small intestine.35
The nature of the immune response to Cyclospora is not clear. Plasma cells have been observed in
lamina propria, and patient sera have demonstrated Cyclospora-specific antibodies. The extraintestinal
Biology of Foodborne Parasites
complications such as biliary disease (cholangitis), acalculous cholecystitis, Guillain–Barre syndrome
or acute febrile polyneuritis, Reiter syndrome, pulmonary infection, and low hemoglobin concentration in the absence of efficient immune system have been recorded in some patients with chronic
Cyclospora infection.36–38
6.7 Treatment and Prevention
The treatment of choice for Cyclospora infections is trimethoprim–sulfamethoxazole (TMP–SMX). It
is reported to be effective for the treatment of Cyclospora infection in both immunocompetent and
immunocompromised patients. In HIV-infected patients, the high rate of recurrence can be reduced by a
prolonged therapy with TMP–SMX. Current regiments of treatment are as follows:
1. For children: Three-day treatment with TMP–SMX at 5–25 mg/kg of body weight/day.39,40
2. For immunocompetent adults: TMP 160 mg plus SMX 800 mg (one double-strength tablet),
orally, twice a day, for 7 days.
3. HIV-infected patients: Longer courses of therapy of 10 days are needed as the relapse rate is
high (43%). Maintenance therapy with double-strength TMP–SMX three times per week prevents relapses.41
Although not as effective as TMP–SMX, Ciprofloxacin is recommended at 500 mg, orally twice a day
for 7 days for patients with sulfa drug allergy.42 Metronidazole, norfloxacin, quinacrine, nalidixic acid,
tinidazole, and diloxanide furoate have also been used in various trials, without success.35,38,43 In Nepal,
a metronidazole and co-trimoxazole combination therapy was effective against cyclosporiasis.44
Since infection occurs most likely by ingestion of feces-contaminated food and water in endemic locations, a few simple solutions are recommended to prevent C. cayetanensis infection.
Travelers should be aware that certain locations are endemic for Cyclospora (generally tropical and
subtropical regions such as Peru, Brazil, Haiti, and Indian subcontinent), especially during the high season for infection when it can more likely be acquired. Avoid ingesting food or water that is not sanitized
properly, and keep proper hygienic habits, food washing, and sanitization.
Strategies to prevent foodborne and waterborne contamination should be targeted for the reduction or
prevention of human infections45:
1. Good agricultural practices to reduce the burden of parasite contamination at the farm level.
These practices would involve the use of properly treated irrigation water and the use of
­pathogen-free water for washing.
2. The safety of local and imported foods needs to be monitored.
3. Adequate processing of water and foods and abstaining from consumption of raw products
when traveling to areas of endemicity.
4. Chemoprophylaxis with TMP–SMX is usually indicated in immunocompromised patients.
A study reported the use of ozone to inactivate Cyclospora oocysts in water.46 More studies are needed
to support this claim. Oocysts survive in chlorinated water but are unlikely to survive the temperature
achieved in anaerobic digestion and do not survive well under low-moisture conditions.47 Cyclospora
cannot yet be propagated in vitro or in vivo; therefore, research has been done using surrogate organisms to identify control strategies effective at killing Cyclospora. One of such studies used Toxoplasma
gondii, another coccidia of public health relevance. Unsporulated oocysts were rendered noninfectious
using 137Cs gamma irradiation at 0.5 kGy on fruits and vegetables, and it was proposed that this treatment
would work similarly with Cyclospora oocysts.48
Cyclospora cayetanensis
6.8 Conclusions
Cyclospora continues to cause outbreaks of diarrheal illness in endemic regions of the world as well as
in nonendemic developed countries that have imported foods contaminated with Cyclospora oocysts.
Trace-back tools and intervention strategies to kill or remove the oocysts from contaminated fresh foods
are not available. Thermal treatment can kill this parasite. The implementation of good agricultural
practices and good food handling and processing practices can reduce the risk of acquiring the infection
not only in the developed countries but also in the countries where these food items are produced for
local consumption.
1. Sun, T. et al. Light and electron microscopic identification of Cyclospora species in the small intestine. Evidence of the presence of asexual life cycle in human host. Am. J. Clin. Pathol. 105, 216–220
2. Ortega, Y.R., Gilman, R.H., and Sterling, C.R. A new coccidian parasite (Apicomplexa: Eimeriidae) from
humans. J. Parasitol. 80, 625–629 (1994).
3. Ortega, Y.R. et al. Pathologic and clinical findings in patients with cyclosporiasis and a description of
intracellular parasite life-cycle stages. J. Infect. Dis. 176, 1584–1589 (1997).
4. Nhieu, J.T. et al. Identification of intracellular stages of Cyclospora species by light microscopy of thick
sections using hematoxylin. Human Pathol. 27, 1107–1109 (1996).
5. Zhao, G.H. et al. Molecular characterization of Cyclospora-like organisms from golden snub-nosed monkeys in Qinling Mountain in Shaanxi province, northwestern China. PLoS ONE 8, e58216 (2013).
5a. Eberhard, M.L., da Silva, A.J., Lilley, B.G., and Pieniazek, N.J. Morphologic and molecular characterization of new Cyclospora species from Ethiopian monkeys: C. cercopitheci, sp.n., C. colobi sp.n., and
C. papionis sp.n. Emerg. Infect. Dis. 5, 651–658 (1999).
6. Zerpa, R., Uchima, N., and Huicho, L. Cyclospora cayetanensis associated with watery diarrhoea in
Peruvian patients. J. Trop. Med. Hyg. 98, 325–329 (1995).
7. Garcia-Lopez, H.L., Rodriguez-Tovar, L.E., and Medina-De la Garza, C.E. Identification of Cyclospora
in poultry. Emerg. Infect. Dis. 2, 356–357 (1996).
8. Yai, L.E., Bauab, A.R., Hirschfeld, M.P., de Oliveira, M.L., and Damaceno, J.T. The first two cases of
Cyclospora in dogs, Sao Paulo, Brazil. Rev Inst. Med. Trop. Sao Paulo 39, 177–179 (1997).
9. Eberhard, M.L. et al. Attempts to establish experimental Cyclospora cayetanensis infection in laboratory
animals. J. Parasitol. 86, 577–582 (2000).
10. Clarke, S.C. Laboratory diagnosis and autofluorescence of Cyclospora. Br. J. Biomed. Sci. 52, 231–232
11. Visvesvara, G.S., Moura, H., Kovacs-Nace, E., Wallace, S., and Eberhard, M.L. Uniform staining of
Cyclospora oocysts in fecal smears by a modified safranin technique with microwave heating. J. Clin.
Microbiol. 35, 730–733 (1997).
12. Galvan-Diaz, A.L., Herrera-Jaramillo, V., Santos-Rodriguez, Z.M., and Gado-Naranjo, M. (Modified
Ziehl–Neelsen and modified Safranin staining for diagnosing Cyclospora cayetanensis). Rev. Salud
Publica (Bogota) 10, 488–493 (2008).
13. Ortega, Y.R. and Sanchez, R. Update on Cyclospora cayetanensis, a food-borne and waterborne parasite.
Clin. Microbiol. Rev. 23, 218–234 (2010).
14. Ortega, Y.R., Sterling, C.R., Gilman, R.H., Cama, V.A., and Diaz, F. Cyclospora species—A new protozoan pathogen of humans. N. Engl. J. Med. 328, 1308–1312 (1993).
15. Lopez, A.S. et al. Epidemiology of Cyclospora cayetanensis and other intestinal parasites in a community in Haiti. J. Clin. Microbiol. 41, 2047–2054 (2003).
16. Jinneman, K.C. et al. Template preparation for PCR and RFLP of amplification products for the detection
and identification of Cyclospora sp. and Eimeria spp. oocysts directly from raspberries. J. Food Prot. 61,
1497–1503 (1998).
Biology of Foodborne Parasites
17. Jinneman, K.C. et al. An oligonucleotide-ligation assay for the differentiation between Cyclospora and
Eimeria spp. polymerase chain reaction amplification products. J. Food Prot. 62, 682–685 (1999).
18. Mundaca, C.C. et al. Use of PCR to improve diagnostic yield in an outbreak of cyclosporiasis in Lima,
Peru. Trans. Roy. Soc. Trop. Med. Hyg. 102, 712–717 (2008).
19. Torres-Slimming, P.A. et al. Outbreak of cyclosporiasis at a naval base in Lima, Peru. Am. J. Trop. Med.
Hyg. 75, 546–548 (2006).
20. Shields, J.M. and Olson, B.H. PCR-restriction fragment length polymorphism method for detection of
Cyclospora cayetanensis in environmental waters without microscopic confirmation. Appl. Environ.
Microbiol. 69, 4662–4669 (2003).
21. Adam, R.D., Ortega, Y.R., Gilman, R.H., and Sterling, C.R. Intervening transcribed spacer region 1 variability in Cyclospora cayetanensis. J. Clin. Microbiol. 38, 2339–2343 (2000).
22. Sulaiman, I.M., Torres, P., Simpson, S., Kerdahi, K., and Ortega, Y. Sequence characterization of heat
shock protein gene of Cyclospora cayetanensis isolates from Nepal, Mexico, and Peru. J. Parasitol. 99,
379–382 (2013).
23. Ortega, Y.R. et al. Isolation of Cryptosporidium parvum and Cyclospora cayetanensis from vegetables
collected in markets of an endemic region in Peru. Am. J. Trop. Med. Hyg. 57, 683–686 (1997).
24. Shields, J.M., Lee, M.M., and Murphy, H.R. Use of a common laboratory glassware detergent improves
recovery of Cryptosporidium parvum and Cyclospora cayetanensis from lettuce, herbs and raspberries.
Int. J. Food Microbiol. 153, 123–128 (2012).
25. Robertson, L.J. and Gjerde, B. Occurrence of parasites on fruits and vegetables in Norway. J. Food Prot.
64, 1793–1798 (2001).
26. Robertson, L.J., Gjerde, B., and Campbell, A.T. Isolation of Cyclospora oocysts from fruits and vegetables using lectin-coated paramagnetic beads. J. Food Prot. 63, 1410–1414 (2000).
27. Shields, J.M., Joo, J., Kim, R., and Murphy, H.R. Assessment of three commercial DNA extraction kits
and a laboratory-developed method for detecting Cryptosporidium and Cyclospora in raspberry wash,
basil wash and pesto. J. Microbiol. Methods 92, 51–58 (2013).
28. Zhou, Y. et al. Prevalence and molecular characterization of Cyclospora cayetanensis, Henan, China.
Emerg. Infect. Dis. 17, 1887–1890 (2011).
29. Herwaldt, B.L. Cyclospora cayetanensis: A review, focusing on the outbreaks of cyclosporiasis in the
1990s. Clin. Infect. Dis. 31, 1040–1057 (2000).
30. Dixon, B., Parrington, L., Cook, A., Pollari, F., and Farber, J. Detection of Cyclospora, Cryptosporidium,
and Giardia in ready-to-eat packaged leafy greens in Ontario, Canada. J. Food Prot. 76, 307–313 (2013).
31. Kozak, G.K., Macdonald, D., Landry, L., and Farber, J.M. Foodborne outbreaks in Canada linked to
produce: 2001 through 2009. J. Food Prot. 76, 173–183 (2013).
32. Kasper, M.R. et al. Diarrhea outbreak during U.S. military training in El Salvador. PLoS ONE 7, e40404
33. Rabold, J.G. et al. Cyclospora outbreak associated with chlorinated drinking water. Lancet 344, 1360–
1361 (1994).
34. Gibbs, R.A. et al. An outbreak of Cyclospora infection on a cruise ship. Epidemiol. Infect. 141, 508–516
35. Connor, B.A., Reidy, J., and Soave, R. Cyclosporiasis: Clinical and histopathologic correlates. Clin.
Infect. Dis. 28, 1216–1222 (1999).
36. Connor, B.A., Johnson, E.J., and Soave, R. Reiter syndrome following protracted symptoms of
Cyclospora infection. Emerg. Infect. Dis. 7, 453–454 (2001).
37. Richardson, R.F., Jr., Remler, B.F., Katirji, B., and Murad, M.H. Guillain-Barre syndrome after
Cyclospora infection. Muscle Nerve 21, 669–671 (1998).
38. Murray, P., Rosenthal, K., and Pfaller, M. In Medical Microbiology (Murray, P., Rosenthal, K., and
Pfaller, M., eds.), pp. 847–860 (Elsevier Mosby, Philadelphia, PA, 2005).
39. Madico, G., Gilman, R.H., Miranda, E., Cabrera, L., and Sterling, C.R. Treatment of Cyclospora infections with co-trimoxazole. Lancet 342, 122–123 (1993).
40. Madico, G., McDonald, J., Gilman, R.H., Cabrera, L., and Sterling, C.R. Epidemiology and treatment of
Cyclospora cayetanensis infection in Peruvian children. Clin. Infect. Dis. 24, 977–981 (1997).
41. Pape, J.W., Verdier, R.I., Boncy, M., Boncy, J., and Johnson, W.D., Jr. Cyclospora infection in adults infected
with HIV. Clinical manifestations, treatment, and prophylaxis. Ann. Intern. Med. 121, 654–657 (1994).
Cyclospora cayetanensis
42. Verdier, R.I., Fitzgerald, D.W., Johnson, W.D., Jr., and Pape, J.W. Trimethoprim-sulfamethoxazole compared with ciprofloxacin for treatment and prophylaxis of Isospora belli and Cyclospora cayetanensis infection in HIV-infected patients. A randomized, controlled trial. Ann. Intern. Med. 132, 885–888
43. Shlim, D.R. et al. An alga-like organism associated with an outbreak of prolonged diarrhea among foreigners in Nepal. Am. J. Trop. Med. Hyg. 45, 383–389 (1991).
44. Sherchand, J.B. and Cross, J.H. Parasitic Epidemiological studies of Cyclospora cayetanensis in Nepal.
Southeast Asian J. Trop. Med. Pub. Health 35, 1–8 (2004).
45. Erickson, M.C. and Ortega, Y.R. Inactivation of protozoan parasites in food, water, and environmental
systems. J. Food Prot. 69, 2786–2808 (2006).
46. Khalifa, A.M., El Temsahy, M.M., and Abou El Naga, I.F. Effect of ozone on the viability of some protozoa in drinking water. J. Egypt. Soc. Parasitol. 31, 603–616 (2001).
47. Gerba, C.P., Pepper, I.L., and Whitehead, L.F., III. A risk assessment of emerging pathogens of concern
in the land application of biosolids. Water Sci. Technol. 46, 225–230 (2002).
48. Dubey, J.P., Thayer, D.W., Speer, C.A., and Shen, S.K. Effect of gamma irradiation on unsporulated and
sporulated Toxoplasma gondii oocysts. Int. J. Parasitol. 28, 369–375 (1998).
49. Huang, P., Weber, J.T., Sosin, D.M., Griffin, P.M., Long, E.G., and Murphy, J.J. The first reported
­outbreak of diarrheal illness associated with Cyclospora in the United States. Ann. Intern. Med. 123,
409–414 (1995).
50. Aksoy, U. et al. First reported waterborne outbreak of cryptosporidiosis with Cyclospora co-infection in
Turkey. Euro Surveill. 12, E070215 (2007).
51. Fleming, C.A., Caron, D., Gunn, J.E., and Barry, M.A. A foodborne outbreak of Cyclospora cayetanensis
at a wedding: Clinical features and risk factors for illness. Arch. Intern. Med. 158, 1121–1125 (1998)
52. Herwaldt, B.L. and Ackers, M.L. An outbreak in 1996 of cyclosporiasis associated with imported raspberries. The Cyclospora Working Group. N. Engl. J. Med. 336, 1548–1556 (1997).
53. Herwaldt, B.L. and Beach, M.J. The return of Cyclospora in 1997: Another outbreak of cyclosporiasis in
North America associated with imported raspberries. The Cyclospora Working Group. Ann. Intern. Med.
130, 210–220 (1999).
54. CDC. Update: Outbreaks of cyclosporiasis- United States and Canada. MMWR 46, 521–523 (1997).
55. CDC. Outbreak of Cyclosporiasis—Ontario, Canada, May 1998. MMWR 47, 806–809 (1998).
56. Ho, A.Y. et al. Outbreak of cyclosporiasis associated with imported raspberries, Philadelphia,
Pennsylvania, 2000. Emerg. Infect. Dis. 8, 783–788 (2002).
57. Botero-Garces, J., Montoya-Palacio, M.N., Barguil, J.I., and Castano-Gonzalez, A. An outbreak of
Cyclospora cayetanensis in Medellin, Colombia. Rev. Salud Publica (Bogota) 8, 258–268 (2006).
58. CDC. Outbreak of Cyclosporiasis—Northern Virginia-Washington, D.C.-Baltimore, Maryland,
Metropolitan Area, 1997. MMWR 46, 689–691 (1997).
59. Lopez, A.S. et al. Outbreak of cyclosporiasis associated with basil in Missouri in 1999. Clin. Infect. Dis.
32, 1010–1017 (2001).
60. Hoang, L.M. et al. Outbreak of cyclosporiasis in British Columbia associated with imported Thai basil.
Epidemiol. Infect. 133, 23–27 (2005).
61. Hammond, R. Cyclospora outbreak in Florida, 2005, abstr. S15. Foodborne Threats Health Policies
Pract. Surveill. Prev. Outbreak Invest. Int. Coord. Workshop, Washington, DC, October 25–26, 2005
62. Doller, P.C. et al. Cyclosporiasis outbreak in Germany associated with the consumption of salad. Emerg.
Infect. Dis. 8, 992–994 (2002).
63. Ayala-Gaytan, J.J., Diaz-Olachea, C., Riojas-Montalvo, P., and Palacios-Martinez, C. Cyclosporidiosis:
Clinical and diagnostic characteristics of an epidemic outbreak. Rev. Gastroenterol. Mex. 69, 226–229
64. CDC. Outbreak of cyclosporiasis associated with snow peas—Pennsylvania. MMWR 53, 876–878
65. CDC. Outbreaks of Pseudo-Infection with Cyclospora and Cryptosporidium—Florida and New York
City. MMWR 46, 354–358 (1995).
66. Blans, M.C., Ridwan, B.U., Verweij, J.J., Rozenberg-Arska, M., and Verhoef, J. Cyclosporiasis outbreak,
Indonesia. Emerg. Infect. Dis. 11, 1453–1455 (2005).
Biology of Foodborne Parasites
67. Ozdamar, M.T., Turkoglu, S., and Hakko, E. Outbreak of cyclosporiasis in Istanbul, Turkey, during an
extremely dry and warm summer, p.2008 988. Abstr. 18th Cong. Clin. Microbiol. Infect. Dis., Barcelona,
Spain (2008).
68. CNEWS. ‘Chef’s Challenge sickens 190 Ontarians—Canoe News
69. Foodborne Outbreak Online Database (FOOD)
70. CDC. Foodborne outbreaks of cyclosporiasis in Texas and Illinois, February 2004. CDC 25133-DS1.pdf
May 24, 2004.
71. WSB-TV. Foodborne illness sickens 100+ at Georgia Aquarium.
72. CDC. Cyclosporiasis outbreak investigations—United States, 2013 (Final Update)
Jorge Néstor Velásquez and Silvana Carnevale
Morphology and Classification......................................................................................................112
7.2.1 Morphology.......................................................................................................................112
7.2.2 Classification.....................................................................................................................114
7.3 Biology, Genetics, and Genomics..................................................................................................114
7.3.1 Biology...............................................................................................................................114
7.3.2 Genetics and Genomics.....................................................................................................115
7.4 Diagnosis and Typing.....................................................................................................................116
7.4.1 Microscopy........................................................................................................................116
7.4.2 Molecular Diagnosis of C. belli........................................................................................119
7.4.3 Typing............................................................................................................................... 120
7.4.4 Molecular Identification of Cystoisospora spp. Other Than C. belli............................... 122
7.5 Epidemiology and Molecular Epidemiology................................................................................ 122
7.6 Pathogenesis and Clinical Features.............................................................................................. 123
7.6.1 Pathogenesis..................................................................................................................... 123
7.6.2 Clinical Manifestations in Humans.................................................................................. 124
7.6.3 Clinical Manifestations in Mammals............................................................................... 126
7.7 Treatment and Prevention............................................................................................................. 126
7.7.1 Treatment.......................................................................................................................... 126
7.7.2 Prevention......................................................................................................................... 127
References............................................................................................................................................... 127
7.1 Introduction
The phylum Apicomplexa contains many organisms of veterinary and medical importance. Coccidia
make up a large portion of this phylum. The coccidia that cause human and animal infections include
Toxoplasma gondii, Sarcocystis spp., Cyclospora spp., and Isospora spp.1 The genus Isospora forms
oocysts with two sporocysts, containing four sporozoites each. They are intracellular parasites, mainly
of the gastrointestinal tract of the host. The life cycle consists of three stages: merogony (asexual cycle),
gametogony (sexual cycle) both taking place in the host, and finally sporogony, which takes place outside
the host. About 200 species of Isospora have been named.2
Morphological and molecular characterizations now differentiate the Isospora-type coccidia into
two apparently monophyletic groups of parasites, the Isospora (Eimeriidae) and Cystoisospora
(Sarcocystidae).3,4 Isospora includes coccidia with tetrasporozoic, diplosporocystic oocysts possessing
Biology of Foodborne Parasites
sporocysts with stiedae bodies, whereas Cystoisospora includes coccidia with tetrasporozoic, diplosporocystic oocysts with sporocysts lacking stiedae bodies.3,4
Cystoisospora belli is coccidia responsible for human cystoisosporosis.2 At present, C. belli infects
human but not other hosts,3 although in one study, infections were reported in gibbons. In
immunocompetent individuals, infection may be asymptomatic or with a moderate to severe illness characterized by fever, vomiting, diarrhea, and abdominal pain.5 In patients with acquired
immunodeficiency syndrome (AIDS), it has been described as another opportunistic agent that
can cause chronic diarrhea, acalculous cholecystitis, and cholangiopathy.6,7 Reports of disseminated cystoisosporosis with unizoite tissue cysts in the lamina propria of the intestines, lymph
nodes, liver, and spleen in patients with AIDS have been published.8–12
Cystoisospora felis infects cats and several other hosts.13–15 C. felis has an extraintestinal cycle
with unizoite tissue cysts in cats as definitive hosts and in other hosts.13–15 In cats, infection may
be asymptomatic or cause enteritis, emaciation, and death.2
Cystoisospora rivolta infects cats, causing disease in newborn kittens, and has extraintestinal
stages in the feline definitive host and in other paratenic hosts.2 Another species, Cystoisospora
ohioensis, infects dogs and other hosts.16,17 C. ohioensis has an extraintestinal cycle with
unizoite tissue cysts in dogs as definitive hosts and in other hosts such as mice and broiler
chicken.13,16,17 It can cause diarrhea.2
Cystoisospora canis infects dogs, has extraintestinal stages in the dog definitive host and in other
hosts,18 and has been shown to be the primary cause of severe diarrhea in 8-week-old female
beagle pups.19 Cystoisospora burrowsi and C. neorivolta also can be found in dog feces.2
Cystoisospora suis is the cause of neonatal porcine coccidiosis.20 Infected piglets develop diarrhea.20 Extraintestinal stages of C. suis have not been described in tissues from infected piglets
or in experimentally inoculated mice.21,22
7.2 Morphology and Classification
7.2.1 Morphology
Oocyst structure. Oocysts that are passed in the feces are ovoid or ellipsoidal. Three distinct layers occur
in the oocyst walls of C. canis and I. canaria and I. serini. The inner, middle, and outer layers are electron-lucent, dense, and lucent, respectively. The oocyst has a round structure inside called a sporont2,23
and is considered not sporulated. The sporont divides to form two uninucleate sporoblasts. The oocyst
is ovoid or ellipsoidal with two round structures inside. A few oocysts that are passed in feces are in the
sporoblast stage.2,23 The sporoblasts elongate and form the sporocyst stage. The genus Cystoisospora
includes oocysts with sporocysts that lack stiedae bodies.3,4 When the sporozoites are fully visible, the
oocyst is considered to be sporulated.
Caryospora-like oocysts contain eight sporozoites in one sporocyst.23 The development of this stage
occurs in approximately 2%–5% oocysts during sporogony in a range of 5 days to 2 weeks at ambient
temperature of approximately 25°C–30°C.2,24,25 Caryospora-like oocysts have been reported in several
species including C. belli, C. canis, C. suis, and C. rivolta.2
Coccidia are identified to the species level based on the structure of their sporulated oocysts. The size,
shape, color, texture, and type of internal contents are important features used in identifying coccidial
oocysts (Table 7.1).
Sporozoite structure. The sporozoites of mammalian Cystoisospora species are elongated and
banana shaped.26 The sporozoites are bounded by an outer membrane unit and an inner membrane complex that apparently consists of two contiguous membrane units.26 At the anterior
end, the sporozoite has an apical complex with conoid, micronemes, rhoptries, polar ring,
and duct-like structures.26 The sporozoite has a single nucleus and contains one or two inclusions named crystalloid bodies.2,26 The crystalloid bodies have no surrounding membrane and
Oocyst Characteristics and Sporulation Times for Cystoisospora Species from Mammals
Cystoisospora Species
C. belli
C. canis
C. ohioensis
C. burrowsi
C. neorivolta
C. rivolta
C. felis
C. suis
Dimensions of Oocysts (μm)
Sporulation Time (Hours)
23–36 × 12–17
34–40 × 28–32
19–27 × 18–23
17–22 × 16–19
18–28 × 16–23
38–51 × 27–39
17–25 × 16–21
Spherical to ovoid
Notes: ?, not described; ??, oocysts are considered to be of the same size as C. ohioensis but were not described.
consist of spherical subunits of nearly uniform size.26 They are composed of particles similar
in appearance to beta-glycogen particles.2 These inclusions are generally lost in the development from sporozoite to merozoite stage in vivo.2 The crystalloid bodies have been identified in
sporozoites of C. belli, C. canis, C. felis, and C. ohioensis.27,28
Merozoite structure. The merozoites are situated within a parasitophorous vacuole in the host epithelial cell. The parasitophorous vacuole contains three membrane units in C. felis and C. rivolta. Each merozoite is invested with an outer membrane unit and an inner double-membrane
complex. The merozoite has an anterior conoid, micronemes, and rhoptries.29 The merozoite
can have a single nucleus, or it can be binucleated or multinucleated.2 Other organelles include
perinuclear Golgi apparatuses, ribosomes, endoplasmic reticulum, mitochondria, and electronlucent and electrodense bodies or granules.29 The merozoite has no crystalloid body.2 The location and site in the intestines and the number of asexual generations vary with each species of
Macrogamete structure. The macrogametes are situated in a parasitophorous vacuole in the host
epithelial cell.2 The membrane of the parasitophorous vacuole of C. felis and C. rivolta has
deep finger-shaped evaginations into the host cell. The macrogamete has a three-layered membrane with a single nucleus and a prominent nucleolus.2 The mitochondria are of the “tubular” type. The macrogametes have electron-lucent and electrodense bodies or granules.2 The
electron-lucent bodies or granules are polysaccharides. The electrodense bodies or granules are
wall-forming bodies of types I and II walls. Type I wall–forming bodies are homogeneous, limited by a membrane unit. Type II wall–forming bodies have no limiting membrane but remain
surrounded by a membrane of the rough endoplasmic reticulum.30 Other organelles include
perinuclear Golgi apparatuses, ribosomes, and endoplasmic reticulum.30
Microgamont and microgamete structure. The microgamont is situated within a parasitophorous
vacuole, which is limited by a multimembranous wall.31 The gamont is enclosed by a membrane
unit. The initial phase of microgamont development consists of cytoplasm growth accompanied by a number of nuclear divisions. The nuclei are initially spheroidal;31 but in later stages
of development, the nuclei migrate to the peripheral zone of the microgamont.31 Each nucleus
contains peripherally condensed chromatin. After completion of the nuclear division stage, the
differentiation of the microgametes begins. The mature microgamete consists of an elongate
nucleus with an elongated mitochondrion and two basal bodies with attached flagella.2
Zygote structure. The macrogamete once fertilized by a microgamete develops into a zygote.30
Type I wall–forming bodies relocate to the periphery of the parasite, disaggregate, and
appear to fuse together at the surface of the parasite forming the outer layer of the oocyst
wall. After the type I wall–forming bodies form the outer layer, type II wall–forming bodies
located in the rough endoplasmic reticulum are transferred via the Golgi body and secretory
granules to the surface where they fuse to create the inner layer of the oocyst wall.30 After
completion of this process, wall-forming bodies are no longer present in the young zygote.30
Biology of Foodborne Parasites
Zygotes contain a central and homogeneous nucleus. The cytoplasm contains numerous amylopectin bodies, canaliculi, and a few lipid vacuoles.30 After maturation, the zygotes form a
wall and develop to oocysts.
Unizoite tissue cyst structure. The unizoite or zoite is located within a large parasitophorous
vacuole and is lined by a three-layered wall.16,28 The parasitophorous vacuole surrounds
the zoite. The parasitophorous vacuole is in close association with the outer portion of the
granular material.27,28 This vacuole is limited by a membrane unit.2 Numerous granules are
filed along the limiting membrane of the parasitophorous vacuole.2,28 The nucleus is situated in the anterior part of the parasite, and the posterior part contains a crystalloid body.
The nucleus is usually ovoid or spherical. The crystalloid body is composed of electrodense
spherical granules.2,28 The crystalloid body has a similar appearance to beta-glycogen particles.2,28 The micronemes are observed predominantly at the apical and posterior poles.
The cytoplasm also contains free ribosomes, Golgi apparatus, endoplasmic reticulum, and
amylopectin bodies.2,28
7.2.2 Classification
Isospora belongs to kingdom Protozoa, phylum Apicomplexa, class Coccidia. The traditional classification of coccidia was based on the number of sporocysts per oocyst and the number of sporozoites
per sporocyst. Two sporocysts, each with four sporozoites, comprise of medically and veterinary
important genera as Besnoitia, Hammondia, Isospora, Neospora, Sarcocystis, and Toxoplasma.2
The traditional classification of coccidian, however, has been repeatedly questioned.3 The phylogenetic positions of members of the Eimeriidae and Sarcocystidae based on the small subunit ribosomal RNA (SSU rRNA) gene (which has been shown to provide good phylogenetic resolution at
the generic and specific levels) have been widely used in diagnostics and taxonomy.32,33 The genus
Isospora was found to be polyphyletic, consisting of two distantly related clades. The Isospora species that contained the stieda and substieda bodies are avian parasites that showed close affinity
to the family Eimeriidae and were assigned to the genus Isospora.4,34 The Isospora species from
mammalian hosts that possessed sutures in the sporocyst wall and lacked the stieda and substieda
bodies, grouped together within the Sarcocystidae, were assigned to the genus Cystoisospora.4,34
Phylogenetic analysis of the internal transcribed spacer 1 of the rRNA (ITS1 rRNA) sequences
of C. felis, C. rivolta, C. ohioensis like, C. belli, C. suis, T. gondii, N. caninum, Sarcocystis, and
Eimeria spp. using distance, minimum evolution, and parsimony-based methods showed that the
genus Cystoisospora does not belong to the family Eimeriidae but should be classified in the family
7.3 Biology, Genetics, and Genomics
7.3.1 Biology
The life cycle consists of endogenous and exogenous development. The endogenous development includes
an asexual cycle or merogony, a sexual cycle or gametogony, and extraintestinal stages that take place in
the host. Sporogony is exogenous and takes place outside the host.
Upon ingestion of sporulated oocysts by a suitable host, sporozoites are released and invade cells of the
intestines of the host. The location and site in the intestines vary by species of Cystoisospora (Table 7.2).
In the asexual cycle or merogony, the sporozoites form meronts and multiply by endodyogeny to form
two daughter merozoites.36,37 The merozoites multiply further by endodyogeny for an indefinite number
of times.2 In the sexual cycle or gametogony, the last generation of merozoites becomes gamonts inside
new host cells and develop either into macrogamonts (female gamonts) or microgamonts (male gamonts).
Biologic Characteristics of Cystoisospora Species from Mammals
Period (Days)
C. belli
C. canis
Period (Days)
C. ohioensis
C. burrowsi
C. neorivolta
C. rivolta
C. felis
C. suis
Development Site
Epithelial cells of small
Cells in lamina propria of small
Epithelial cells of small
intestine, cecum, and colon
Epithelial cells and cells in
lamina propria of small
Cells in lamina propria
Epithelial cells of small
Epithelial cells of small
intestine, occasionally cecum
Epithelial cells of small
intestine, cecum, and colon
Types of Meronts
Types I, II, III
Types I, II
(uni-, bi-, multinucleated)
Types I, II
(not multinucleated)
Types I, II
(uni- or binucleated), III, IV
Types I, II
(multinucleated), III
Types I, II
(uni- or multinucleated), III
Types I (binucleated),
II (multinucleated)
Note: ND, not described.
Microgamonts produce numerous microgametes.2 The microgamete penetrates the macrogamete and
forms the zygote,30 which after maturation develops into an oocyst. The oocysts are released into the
lumen of the intestines and passed via feces unsporulated.
The prepatent and patent periods vary by Cystoisospora species. Sporozoites have also been known to
leave the intestinal tract and infect other tissues in the definitive host and in other hosts. Extraintestinal
stages have been described in the life cycle of C. belli, C. felis, C. rivolta, C. ohioensis, and C. canis
but not in C. suis.2,21,22 If no replication occurs in the intermediate host, it is referred to as a paratenic
host.31 These stages are usually found to be single organisms resembling sporozoites or merozoites in the
intestinal lamina propria, lymph nodes, spleen, and liver.2 C. belli infects humans but not other hosts.2
C. felis has an extraintestinal cycle with unizoite tissue cysts in cats as definitive hosts and in other hosts
including mice, rats, hamsters, birds, rabbit, and swine.13–15 C. ohioensis has an extraintestinal cycle with
unizoite tissue cysts in dogs as definitive hosts and in other hosts such as mice and broiler chickens.13,16,17
C. canis infects dogs and has extraintestinal stages in the definitive host (dogs) and in other hosts including pigs, buffalo, and camels.18
Sporogony takes place outside the host, and oocyst sporulation depends on environmental conditions
such as oxygen, temperature, and humidity.2 Upon sporulation, the sporocysts contain the infective sporozoites.2 The sporulation period varies depending on the Cystoisospora species.2,31 Cystoisosporosis
can occur if a definitive host ingests a paratenic host harboring unizoite tissue cysts.2,31
The life cycle of C. belli is shown in Figure 7.1.
7.3.2 Genetics and Genomics
Cystoisospora genomes have not been sequenced, and only 69 Cystoisospora nucleotide sequences are
available in GenBank. All are rRNA sequences, with exception of a partial sequence of the cytochrome
c oxidase subunit 1 (CO1) gene. Out of the 69 sequences, 44 were from C. belli (all rRNA sequences),
8 from C. ohioensis, 6 from C. felis (including the 1 CO1 gene sequence), 5 from C. suis, 2 from
C. ­rivolta, 2 from C. timoni (from suricatas), and 2 from Cystoisospora spp. in dogs, cats, and raccoon
dogs in Japan.
Biology of Foodborne Parasites
Sporulated oocysts
Outside the host
Inside the host
Excystation in
intestinal lumen
Zygote Macrogamont
Lamina propria
Lymph nodes
Extraintestinal stage
FIGURE 7.1 Life cycle of C. belli.
7.4 Diagnosis and Typing
7.4.1 Microscopy
Diagnosis of cystoisosporosis is based on the identification of the oocyst stage of the parasite in fecal
samples. Assays may be performed with or without prior fixation, depending on the method. Specimens
should be kept refrigerated if unfixed and sent to the laboratory as promptly as possible. If this is not
possible, then the samples should be preserved. The more commonly used fixatives are 10% neutral
formalin or sodium acetate–acetic acid–formalin,38 but they should not be used if the samples will be
analyzed by PCR. Potassium dichromate may be used if oocyst viability or infectivity is to be assessed
and for processing for molecular diagnosis.38 Unfixed stool samples should be frozen and can be used
for molecular diagnosis.
The oocysts are usually excreted in small numbers and intermittently. Therefore, techniques to concentrate oocysts have generally been used as part of routine diagnostic procedures.39 Three stool specimens from three separate days for detection of oocysts provide better sensitivity than only one sample.38
Stool material can be processed either as direct smears or by flotation and sedimentation concentration
methods, which are more sensitive than direct smears.2,40 Sheather sugar flotation and formalin–ether
sedimentation methods are excellent for detecting oocysts.2,41
Oocysts are visible by light microscopy. The oocyst is ovoid or ellipsoidal with a well-defined
oocyst wall (Figure 7.2). The oocyst is passed unsporulated in the stools and has a round structure
inside,2,23 although a few oocysts passed in feces can have two round structures inside.2,23 Fecal examination commonly reveals Charcot–Leyden crystals.42 Blood and leucocytes are rarely observed.43 The
identification of oocyst in wet preparations can be greatly enhanced by using epifluorescence microscopy. Cystoisospora oocysts are autofluorescent under UV fluorescence microscopy. The oocyst wall
10 μm
FIGURE 7.2 Unsporulated oocyst of C. belli observed by light microscopy after ethyl ether concentration (magnification 400×).
autofluoresces blue–violet under UV light at 330–365 nm.44 The disadvantage is that some laboratories
may not have access to fluorescence microscopy. Another consideration is the fact that wet mount preparations cannot be archived as a permanent record. The oocyst wall and sporonts or sporoblasts fluoresce bright yellow when Cystoisospora oocysts are stained using auramine–rhodamine and should be
confirmed by wet mount smear examination or acid-fast staining.41 The use of acid-fast-stained smears
serves as the standard for detecting coccidian oocysts. The oocyst has the typical ellipsoidal shape. The
oocyst wall is highlighted and has bright red sporonts or sporoblasts.45 Some oocysts may appear collapsed, distorted, or without sporonts or sporoblasts. The advantage of this staining method is that it
provides a permanent record.
The different intracellular parasitic stages of C. belli may be found in the intestinal and biliary epithelial cells in biopsy specimens.39,46–48 Stages of C. belli have been identified in biopsy samples of duodenal, jejunal, ileal, biliary, and gallbladder specimens.39,49,50
The detection of any of these stages of development is diagnostic of coccidiosis.2,39 Routine histological staining methods are satisfactory for demonstrating parasite stages.2 Stages are difficult to identify
in samples stained with hematoxylin and eosin because of the similarity of staining of the host epithelial
cells.39 Organisms stain positively with Giemsa, periodic acid–Schiff (PAS), and methenamine silver
stains.39 The small bowel mucosa shows villous atrophy (shortening, blunting, fusion, and clubbing)
with crypt hyperplasia/hypertrophy.47 The lamina propria is infiltrated by eosinophils, lymphocytes,
and plasma cells.39,47 There can be marked dilation of vascular channels.29,39 The epithelial cells may or
may not appear normal as the pathology of the epithelial cells is varied including vacuolization, changes
in the shape of the cells from columnar to cuboidal, and atrophy of microvilli.29,39 In patients infected
with low numbers of parasites, it is necessary to examine multiple serial sections for identifying the
The meront is situated in a well-developed parasitophorous vacuole. The meront stage has an ellipsoidal or spherical shape (Figure 7.3). The mean measure of the meront stage is about 15 μm × 9 μm. The
number of merozoites produced is highly variable from 1 to 10. The merozoites have a length of 11 μm
and a width of 2.5 μm. The nucleus is unique and does not have a crystalloid body.2 The duration and the
number of generations of schizogony in man are not known.2,39
The microgamont is also situated within a parasitophorous vacuole and is spherical in shape.31 The
mean diameter of the microgamont is about 10 μm. The nuclei in young stages are at first spheroidal. The nuclei are multiple with a homogeneous distribution in the cytoplasm.31 In the later stages
Biology of Foodborne Parasites
10 μm
FIGURE 7.3 Meront of C. belli in duodenal biopsy specimen stained with hematoxylin–eosin (magnification 1000×).
of development, the nuclei migrate to the peripheral zone of the microgamont.31 Mature forms have a
residuum and shed a large number of flagellated, comma-shaped microgametes.2,39
The macrogamete is ellipsoidal in shape (Figure 7.4) and measures ~18 μm × 7 μm. The nucleus is
unique with prominent nucleolus. The cytoplasm is uniformly highly vacuolated and contains a large
number of granules of variable size.2,39 The zygote contains a central and homogeneous nucleus. The
zygote, or young oocyst (Figure 7.5), measures ~25 μm × 8 μm. The cytoplasm contains numerous
10 μm
FIGURE 7.4 Macrogamete of C. belli in duodenal biopsy specimen stained with azure II (magnification 1000×).
10 μm
FIGURE 7.5 Young oocyst of C. belli in duodenal biopsy specimen stained with hematoxylin–eosin (magnification 1000×).
10 μm
FIGURE 7.6 Unizoite of C. belli in duodenal biopsy specimen stained with azure II (magnification 1000×).
The unizoites, depending on tissue section, acquire round, oval, and banana-shaped forms (Figure 7.6).
The unizoite tissue cyst infects host cells whose type is difficult to determine. The cyst wall is eosinophilic in hematoxylin and eosin–stained sections and light blue in azure II–stained sections.2,11 The cyst
wall has been seen as PAS positive in one patient8 and not in others.2,10 Zoites are located in the center of
the tissue cyst, surrounded by a clear vacuole. The zoites are uninucleated. The nuclei are rounded and
anteriorly or centrally located.8–12
7.4.2 Molecular Diagnosis of C. belli
Müller et al.51 described for the first time a DNA-based technique for the detection of C. belli infections.
They developed a PCR/southern blot using primer pairs and a hybridization probe based on variable
regions of the SSU rRNA of C. belli. The confirmation of DNA fragments amplified during the first
Biology of Foodborne Parasites
PCR was done either by nested PCR or by southern blot hybridization. The assay was tested in intestinal biopsies and bile from two patients. The nested PCR was negative in samples from patients with
eight different intestinal pathogens (Cryptosporidium parvum, Giardia duodenalis, Blastocystis hominis, Endolimax nana, Entamoeba hartmanni, Entamoeba histolytica, Enterocytozoon bieneusi, and
Encephalitozoon intestinalis) but gave a weak signal at the expected length with template DNA derived
from T. gondii. No other species belonging to the genus Cystoisospora were included in the study.
Samarasinghe et al.35 developed a PCR–RFLP assay based on the ITS1 sequence to detect and differentiate Cystoisospora species. The external primers were designed based on conserved sequences,
and the internal primers were specific for Cystoisospora as they did not amplify DNA from C. parvum,
G. duodenalis, Neospora caninum, T. gondii, Eimeria tenella, E. maxima, E. acervulina, Sarcocystis
canis, and S. felis. RFLP analysis by digestion with AluI enzyme produced distinct banding patterns that
differentiated C. belli, C. suis, C. felis, C. rivolta, and C. ohioensis.
A real-time PCR assay targeting the internal transcribed spacer 2 region of the ribosomal RNA gene
(ITS2) was developed by ten Hove et al.52 for the detection of C. belli DNA in fecal samples, including an internal control to detect PCR inhibition. The specificity of the assay was evaluated using 27
control DNAs from parasites (Schistosoma mansoni, Necator americanus, Strongyloides stercoralis,
Ancylostoma duodenale, Cyclospora cayetanensis, E. histolytica, Entamoeba dispar, G. lamblia, C. parvum, E. bieneusi, E. intestinalis, Dientamoeba fragilis, and T. gondii) and bacteria/yeast (Enterococcus
faecalis, Staphylococcus aureus, Pseudomonas aeruginosa, Salmonella enteritidis, S. typhimurium,
Enterobacter aerogenes, Yersinia enterocolitica, Bacillus cereus, Proteus mirabilis, Shigella flexneri,
S. dysenteriae, S. sonnei, Escherichia coli O157, and Candida albicans,); 80 DNA extracts derived from
feces positive for E. histolytica, E. dispar, G. lamblia, or C. parvum/C. hominis; and 40 DNA extracts
derived from feces of individuals with no known history of parasitic infections and with a negative result
in stool samples by microscopy using formalin–ether sediments and modified acid-fast staining. No
amplification of C. belli–specific DNA was detected in any of these 147 samples. The PCR was evaluated
using DNA extracts of 21 stool samples from individual patients in which the microscopic examination
of modified acid-fast-stained fecal smears revealed C. belli oocysts. C. belli–specific amplification was
detected in all microscopy-positive samples. The assay achieved 100% specificity and sensitivity. The
detection limit was estimated to be one C. belli oocyst.
Taniuchi et al.53 developed a sensitive and specific multiplex PCR reaction with four primer sets to
amplify C. cayetanensis, C. belli, E. bieneusi, and E. intestinalis. Target genes for C. belli included
the 5.8S rRNA and ITS2. Detection of the amplicon was through specific probes coupled to Luminex
beads. The limit of detection was 102 copies of spiked plasmid with Cystoisospora sequence inserts.
No cross-reactivity was observed with positive specimens of Giardia, Cryptosporidium, E. ­histolytica,
Ascaris, Strongyloides, Necator, Ancylostoma, Trichuris, enteroaggregative E. coli, enteroinvasive
E. coli, enteropathogenic E. coli, enterotoxigenic E. coli, enterohemorrhagic E. coli, rotavirus, adenovirus, astrovirus, sapovirus, and norovirus. They evaluated the assay on 236 diarrheal specimens from
Thailand, Tanzania, Indonesia, and the Netherlands. Performance of the multiplex Luminex assay versus
microscopy revealed 87%–100% sensitivity and 88%–100% specificity. For Cystoisospora, the PCR
Luminex assay detected about 50% more infections than microscopy.
Another approach was developed by Murphy et al.54 who employed extended-range molecular screening with widely conserved primer-binding sites in the rRNA genes to detect C. belli infection in a patient
with cystoisosporosis. They included PCR primers capable of detecting a broad range of bacterial, fungal, and some parasitic pathogens, based on the ITS1, ITS2, and the 28S rRNA gene with sequencing of
the amplification products to confirm identification.
7.4.3 Typing
In 1998, Carreno et al.55 showed by phylogenetic analysis that Isospora (employing I. suis, I. ohioensis,
and I. felis) was most closely related to Toxoplasma and Neospora.
Franzen et al.56 sequenced the SSU rRNA of C. belli (at that time still classified as Isospora belli) and
used it for the analysis of the taxonomic position of C. belli. The isolate they used was a bile juice extract
from a patient chronically infected with C. belli. A universal primer pair was used to amplify the entire
SSU rRNA gene of C. belli, and the product was cloned and sequenced. The SSU rRNA gene sequence
of C. belli and those of several members of the phylum Apicomplexa were used for phylogenetic analysis. The phylogenetic trees provided molecular evidence for three clades within a monophyletic group
that represents the suborder Eimeriorina. The clade containing C. belli consists of tissue-cyst-forming
coccidia (Toxoplasma and Neospora) and members of the genus Cystoisospora (C. ohioensis, C. suis,
and C. belli), with these last ones forming a monophyletic group. The second clade, representing a sister
clade of the first one, contains members of the genus Sarcocystis. The third one consists of members
of the family Eimeriidae, including Eimeria and Cyclospora species. This report showed by the first time
that although C. belli and other members of the genus Cystoisospora belong to the suborder Eimeriorina,
the family to which they belong was not Eimeriidae but rather Sarcocystidae.
In 2003, Millar et al.57 sequenced the 5.8S rRNA, ITS2, and 28S rRNA regions from C. belli and used
them to compare the taxonomical position of this species with its closest phylogenetic neighbors, showing that C. belli was most closely related to the genus Besnoitia, followed by N. caninum and T. gondii.
Jongwutiwes et al.5 investigated genetic heterogeneity in Cystoisospora isolates infecting patients in
Thailand. They analyzed 38 fecal samples containing Cystoisospora oocysts from HIV-infected patients
(n = 30), corticosteroid-treated patients (n = 3), and immunocompetent individuals (n = 5). Cystoisosporiasis
was diagnosed by the presence of oocysts in stool samples by direct wet smear method, formalin–­
ethylacetate sedimentation, or acid-fast staining. The morphometry of the oocysts was determined, and
oocysts were sporulated. A DNA fragment spanning the SSU rRNA, ITS1, 5.8S rRNA, and ITS2 regions
(3049 base pairs) of C. belli was amplified by a nested PCR, and the products were cloned and sequenced.
Although oocysts showed shape and size variations both within and between isolates, the shape indices of
all oocysts observed in that study were consistent with that of C. belli. Sequences of the SSU rRNA gene,
spanning 1778 base pairs, of 25 isolates examined from HIV-infected patients were identical with those of
strains CI1 and CJLPHD2 (GenBank accession nos. U94787 and AF441289) but differed from the isolate
reported by Franzen et al.56 (GenBank accession nos. AF106935 at A679T and A682C). Three additional
nucleotide substitutions occurred at T583C, C638A, and G1240T in the isolate from an immunocompetent
patient who had multiple relapses. The 5.8S rRNA and ITS2 regions contained 598 and 404 base pairs,
respectively, and showed perfect sequence identity among all isolates. Sequences of the ITS1 containing 158 base pairs were highly conserved, except for one nucleotide substitution at position 528 with an
A to G change (position according to the ITS1 region). The SSU RNA, 5.8S rRNA, ITS1, and ITS2 were
highly conserved, indicating that there were no cryptic species or extensive strain variation. The neighbor-joining tree derived from the SSU rRNA sequences obtained and those from the genera Toxoplasma,
Cryptosporidium, Cyclospora, Neospora, Eimeria, and Hammondia confirmed that all isolates of C. belli
in that study were clustered, which is consistent with a single species and that C. belli was more related to
Toxoplasma, Neospora, and Hammondia than to Cyclospora, Eimeria, and Cryptosporidium. An identical
topology of the tree was obtained when the ITS2 sequences were used for comparison.
Phylogenetic analysis of ITS1 sequences of Cystoisospora species (C. felis, C. rivolta, C. ohioensis
like, C. belli, and C. suis) was performed by Samarasinghe et al.35 They also employed ITS1 sequences
for Toxoplasma, Neospora, Sarcocystis, and Eimeria spp. using distance, minimum evolution, and
parsimony-based methods. The generated trees, with nearly identical topologies, supported the closer
relationship of Cystoisospora spp. to T. gondii and N. caninum and Sarcocystis species than Eimeria
species. The similarity between different Cystoisospora species ranged from 74.7% to 90.9%. The similarity between C. belli and T. gondii and N. caninum was 76.7% and 75.5%, respectively. The similarity
between C. belli and Sarcocystis species was 69.4%–71.1%, whereas the similarity between C. belli and
Eimeria species was 61.8%–64%. These authors confirmed results of previous studies, which suggested
that the genus Cystoisospora does not belong to the family Eimeriidae but should be classified together
with the cyst-forming coccidia in the family Sarcocystidae.
In 2011, Velásquez et al.58 analyzed samples from eight adult patients from Argentina with AIDS
who were evaluated for chronic diarrhea and cystoisosporosis. They identified Cystoisospora isolates,
with and without the presence of unizoites, with molecular tools based on the SSU rRNA and ITS1
regions. Sequencing of a 396 bp fragment of the SSU rRNA gene did not detect differences between
isolates. Identity was 100% for previously reported sequences for C. belli, with the exception of one
case described by Jongwutiwes et al.5 of an immunocompetent patient with multiple relapses. When
Biology of Foodborne Parasites
compared with other Cystoisospora species, this value reduced to 98.4% but showed high identity values
(over 99%) with the sequences of other Cystoisospora species in which the presence of unizoites in their
life cycles is common, namely, C. ohioensis and C. felis. Sequences of the ITS1 fragment from the eight
isolates were identical between them but differed from the isolates reported by Jongwutiwes et al.5 and
Samarasinghe et al.35 Identity was 99.7% and 99.5% for previously reported C. belli ITS1 sequences and
ranged from 81.9% to 58.8% for other Cystoisospora species. It was not possible to identify species or
strain variation in isolates of C. belli with or without unizoite tissue cysts.
Resende et al.59 analyzed intraspecific variability among 13 C. belli isolates obtained from Brazilian
HIV-infected patients by restriction fragment length polymorphisms (RFLPs) with MboII in a 1.8 kb
amplicon of the SSU rRNA region of the parasite. Three RFLPs were generated, which were named
RFI, RFII, and RFIII. Two isolates obtained from a patient with extraintestinal cystoisosporosis showed
distinct RFLPs with MboII. The authors demonstrated the genetic heterogeneity among clinical isolates
of this parasite, which formed three polymorphic clusters, and that patients can be infected with different C. belli genotypes, supporting the existence of mixed infection with different C. belli genotypes or
specific parasite populations able to invade and multiply in different host tissues.
7.4.4 Molecular Identification of Cystoisospora spp. Other Than C. belli
In 2001, Ruttkowski et al.60 developed a species-specific PCR to differentiate C. suis, Eimeria polita,
E. porci, and E. scabra based on the SSU rRNA region. Johnson et al.61 screened 289 pig fecal samples
in Western Australia for the presence of Cystoisospora sp. using a PCR–RFLP assay based on the ITS1
rRNA locus. They identified an overall prevalence of 10.4% (30/289) and confirmed the presence of
C. suis in 86.7% of the positive Cystoisospora isolates.
Lalonde and Gajadhar62 developed a real-time quantitative PCR assay using melting curve analysis to detect, differentiate, and identify DNA from coccidian species of animal health, zoonotic, and
food safety concern, including C. suis. Matsubayashi et al.63 isolated Cystoisospora spp. from dogs,
cats, and raccoon dogs in Japan and analyzed the isolates by morphology and molecular methods based
on the SSU rRNA gene locus to determine their phylogenetic position. Phylogenetic analysis showed
that the dog and raccoon dog isolates were nested in a clade with other Cystoisospora spp. including
C. ­ohioensis, C. belli, and C. orlovi, while the cat isolate formed a sister group with C. felis. The isolates
from the dogs and raccoon dogs and cats were morphologically and genetically closely related to C. ohioensis and C. felis, respectively, but not completely identical to those of C. ohioensis and C. felis deposited
in GenBank, showing sequence variation.
In 2012, He et al.64 isolated Cystoisospora spp. oocysts from dog feces in China. They showed that
these oocysts were morphologically similar to those of C. ohioensis and Cystoisospora sp. isolated from
dogs in Japan. Phylogenetic analysis of the 18S rRNA sequences showed that the Cystoisospora sp.
isolates from China (Changchun 1 and Changchun 2) were nested in a clade with other Cystoisospora
spp., including C. ohioensis, C. belli, C. suis, Isospora sp. Harbin/01/08, and C. orlovi. Cystoisospora
sp. Changchun 2 was confirmed as C. ohioensis, and the other isolate was relatively close to C. suis after
analysis of the ITS1 gene.
7.5 Epidemiology and Molecular Epidemiology
C. belli infections are essentially cosmopolitan in distribution but are more common in tropical and
subtropical regions.40,43
The prevalence of C. belli in inmunocompetent patients with diarrhea in developed countries with
good economic and sanitary conditions is 1% or less.40,43 Cystoisosporosis is usually found in these countries with high numbers of cases in institutions or schools for mentally challenged, immigrant patients
or people with a history of travel.
In developing countries, the prevalence of C. belli in immunocompetent patients with diarrhea ranges
from <1% to 13.1%.40 In Chile, cases of C. belli were identified in areas with good economic and sanitary
The prevalence of C. belli in AIDS patient with diarrhea in developed countries ranges from <1% to
3%.66 Infection rates are 0.07% in Japan, 1% in the United States (Los Angeles County), and 1%–2% in
Switzerland. Significant risk factors for cystoisosporiasis are foreign-born patients, travel exposure, and
recent immigration.
In developing countries, the prevalences of C. belli in AIDS patient with diarrhea range from 1% to
41%.2,67,68 The prevalences of C. belli in developing countries in pre–highly active antiretroviral therapy
(HAART) era were 15% in Haiti,69 19% in Zaire, and 16% in Zambia. In HAART era, the prevalence can
be low or high: 3.1% in Nigeria, 7.5% in Korea, 9.7% in Egypt, 41% in India,68 12% in Ethiopia, and 10%
in Malaysia. In Brazil, the prevalence of cystoisosporosis in HIV patients ranged from 10% to 18%,67
with differences in the same country. For example, in Ribeirão Preto, the prevalence was 4.4%; in São
Paulo, 5.7%; in Santos, 9.9%; in Rio de Janeiro, 10.1%; and in Campinas, 18%.67
Cystoisosporosis mainly affects young animals.2,70 Cystoisospora suis is one of the most frequent
causative agents in outbreaks of infectious diarrheas in sucking piglets with a cosmopolitan occurrence.71
It is responsible for 17%–54% of the cases of piglet diarrhea seen at diagnostic laboratories, with values
in Northern Europe of 17%, Germany 27%, and Australia 54%. In Australia, C. felis has a prevalence of
1.7%–10% in cats by microscopy.72 The examination of fecal samples from dogs in Germany identified
C. ohioensis complex and C. canis with a prevalence of 3.9% and 2.4%, respectively.73
Fecal screenings for coccidial oocysts have a prevalence of 3%–38% in dogs and 3%–36% in cats.2,70
C. felis has a prevalence of 1.7%–10% in cats in Australia.72 The examination of fecal samples from
dogs revealed C. ohioensis complex and C. canis with a prevalence of 3.9% and 2.4%, respectively, in
Current knowledge on molecular epidemiology of C. belli is mostly based on three reports, all of
which characterized the rRNA locus of the parasite.5,58,59 Samples from Thailand, Brazil, and Argentina
to date have been analyzed, including a total of 47 patients of which 44 were HIV infected and 3 were
immunocompetent. The studies by Franzen et al.56 and Samarasinghe et al.,35 which included sequencing
the rRNA gene or ITS1 region, did not specify the geographical origin of the samples. Some genetic heterogeneity among clinical isolates of this parasite has been shown, but the determination of the extent of
this heterogeneity requires studies with isolates from wider geographic regions. The relationship among
different genotypes of Cystoisospora with the epidemiology of cystoisosporosis, resistance of the parasite to anticoccidial drugs, and clinical manifestations in infected patients is unknown. The use of more
than one genetic marker and more variable markers will be important in identifying genetic variation
within C. belli and improving the understanding of molecular epidemiology.
7.6 Pathogenesis and Clinical Features
7.6.1 Pathogenesis
Acquisition of cystoisosporosis is thought to be through ingestion of food or water contaminated with
mature oocysts from human feces. C. belli oocysts are not immediately infective, so person-to-person
transmission is considered rare.
In immunocompetent individuals, infection may be asymptomatic or with fever, vomiting, diarrhea, abdominal pain, and malabsorption.5 Light microscopy of the small bowel mucosa shows villous atrophy (shortening, blunting, fusion, and clubbing) with crypt hyperplasia/hypertrophy.47 The
lamina propria is infiltrated by eosinophils, lymphocytes, and plasma cells.39,47 In recent years, it has
become apparent that the mucosal immune system plays an important role in crypt proliferation and
villus atrophy.74 Reports showed that during a gastrointestinal infection, T cells are activated.75 Villus
atrophy and crypts were longer with crypt epithelial cell proliferation in T-cell-stimulated cultures.74
In C. suis–infected piglets, the number of a subset of T cells increased in the gut mucosa.76 Infections
with Eimeria bovis, E. falciformis, E. acervulina, E. maxima, and E. tenella stimulate T-cell reactions
during the host response.77–79 Lambs experimentally infected with E. ovinoidalis showed a significant
increase in CD8 lymphocytes in the small intestine epithelium, associated with diarrhea, villous atrophy, and crypt hyperplasia.80
Biology of Foodborne Parasites
The clinical presentations of C. belli in patients with AIDS include acute, persistent, or chronic diarrhea.2 C. belli infection is a cause of crypt hyperplasia and villous atrophy.2 However, crypt hyperplasia
and villous atrophy can occur at all clinical stages of HIV disease and in the absence of a detectable
enteropathogen. The total T-cell population and the component of T helper cells (CD4) are depleted,
while the component of T suppressor cells (CD8) is proportionally increased.81 The mucosal immune
system plays an important role in crypt proliferation and villus atrophy during infection with C. belli
and HIV infection.2,81 These patients can have a number of features that are consistent with malabsorption such as fecal fat, anemia with thrombocytosis (suggesting iron deficiency), an elevated prothrombin
time (suggesting vitamin K deficiency), a low normal serum albumin level (consistent with protein loss
or poor synthesis), and low total blood calcium and phosphorus levels.82 Hypokalemia can be associated
with diarrhea and vomiting.82
7.6.2 Clinical Manifestations in Humans
The clinical presentations of C. belli in patients with AIDS include asymptomatic, symptomatic, and
extraintestinal involvement. There are descriptions of asymptomatic cystoisosporosis in AIDS patients.2
In patients with symptomatic intestinal presentation, clinical symptoms include acute, persistent, or
chronic diarrhea,2 and it is important to take into account the level of CD4 cell counts.83 In patients with
a CD4 count >250 cells/mm3, the diarrhea is acute or persistent.83 Patients with CD4 counts between 100
and 250 cells/mm3 present with chronic diarrhea.83 Patients with CD4 counts <100 cells/mm3 present
with chronic diarrhea and disseminated cystoisosporosis.2 C. belli has been associated with malabsorption syndrome, weight loss, and hypokalemic paralysis.
Extraintestinal symptomatic presentations comprise of acalculous cholecystitis6 and disseminated
cystoisosporosis.8,10,11 In some patients with AIDS, chronic diarrhea, and cystoisosporosis, histological
structures of unizoites were identified in different tissues other than intestinal epithelium.8,10–12 Unizoites
have been described in the lamina propria, lymph nodes, liver, and spleen.8,10–12
The test of patients with AIDS and diarrhea for C. belli begins with the stool examination.84 The stool
test is performed on three or more samples84 and for the diagnosis of cystoisosporosis is advisable at least
four stool samples.85 If the stool analysis is negative, an upper endoscopy with biopsy sampling of the
distal portion of the duodenum is indicated.84
Associated infections with Cryptosporidium sp., G. duodenalis, and Trichuris trichiura were observed
in patients with C. belli identified in feces and a low CD4 cell count.29 Thus, in patients with cystoisosporosis and a CD4 cell count of 100 cells/mm3 or less, it is necessary to evaluate whether or not to
perform upper endoscopy with duodenal biopsy samples to identify Cystoisospora sp. and coinfections.
The examination of biliary tract abnormalities in patients with AIDS consists of ultrasound analysis,
magnetic resonance cholangiography, and endoscopic retrograde cholangiopancreatography. In these
patients, the diagnosis also includes examination of feces and duodenal and papillary biopsy samples.
The clinical manifestations of the six reported patients with disseminated cystoisosporosis are summarized in Table 7.3.
The clinical presentations of C. belli in immunocompetent patients include asymptomatic, symptomatic, and extraintestinal infections with children and travelers more likely to be infected.2 Patients with
a symptomatic intestinal presentation have diarrhea that begins 8 days after oocyst ingestion.69 The
most common symptom is diarrhea (6–10 stools per day), which may be accompanied by abdominal
pain and weight loss.69 In this clinical presentation, patients have persistent or chronic diarrhea.39,47,86
Oocysts are eliminated in the feces usually within 10 or 15 days after infection,2,29 but up to 30 days have
been reported.87 Once diarrhea resolves, untreated patients continue to shed oocysts for 15–30 days.29
Malabsortion, steatorrhea, and eosinophilia are often present in immunocompetent patients.39,88
Extraintestinal symptomatic presentation comprises chronic cholecystitis and sclerosing cholangitis.49 The diagnosis in immunocompetent patients is generally via stool examination.2,29 Examination
of a single stool sample may be negative due to the intermittency in oocyst excretion,89 and in this
case, it is advisable to repeat it.89 The total number of new samples to confirm a negative result has not
been established.2,89 If the stool examination is negative, it is necessary to consider the need for upper
endoscopy with biopsy sampling of the distal portion of the duodenum in patients with persistent or
Yes (during 3 years)
Yes (3 months)
Pneumocystis carinii pneumonia,
G. lamblia, E. histolytica, Mycobacterium
kansasii, cytomegalovirus
Restrepo et al.8
Michiels et al.10
Yes (7 days)
Yes (3 months)
Lymph nodes (mesenteric)
Chronic diarrhea intermittent
208 to 53
Yes (biliary vesicle)
Not described
Yes (1–3 months) Yes (during 5 years)
Cerebral toxoplasmosis,
herpes zoster,
Mycobacterium tuberculosis
Velásquez et al.11 Frenkel et al.12
Yes (4 days)
No adherence
Yes (5 days)
Lamina propria
Chronic diarrhea
Lamina propria
Chronic diarrhea
Comin and Santucci9 Velásquez et al.11
Yes (2 days)
Intravenous drug
Chronic diarrhea, weight loss Chronic diarrhea
Yes (gamont in epithelial
cells of lamina propria)
Lamina propria, lymph
Lamina propria
nodes, liver, spleen
Not described
Lamina propria, lymph nodes
(mesenteric, periaortic, mediastinal)
Vomiting, chronic diarrhea, weight loss
Note: ND, not described.
Unizoite tissue cysts
present in
Positive/negative for
oocyst in stools
Antimicrobial therapy
Antiretroviral therapy
Resolution of
diarrhea (at 1 week)
Other opportunistic
Sex/age (year)
Risk factor for HIV
Asexual stages
Sexual stages
Clinical Manifestations of Patients with Disseminated Cystoisosporosis
Biology of Foodborne Parasites
chronic diarrhea and malabsorption.39 The optimal number of stool samples required to confirm the
absence of C. belli has not been established.2,43
C. belli is a cause of opportunistic infections in patients with other immunodeficiencies including
patients with Hodgkin’s and non-Hodgkin’s lymphomas90 and acute lymphoblastic leukemia91 and in
patients receiving prolonged corticosteroid therapy.92 C. belli infections have also been reported in renal
transplant recipients93 and in a liver transplant patient with chronic diarrhea.94 The clinical presentations
include persistent and chronic diarrhea.90–94
7.6.3 Clinical Manifestations in Mammals
Cystoisospora infections are generally only associated with disease in puppies and kittens. C. canis has
been shown to be the primary cause of severe diarrhea in 8-week-old female beagle pups.19 C. ohioensis
can cause diarrhea in 7-day-old pups but not in weaned pups or young dogs.36 C. felis infection in cats
may be asymptomatic or cause enteritis, emaciation, and death.2 C. rivolta causes disease in newborn
kittens.37 C. suis is the cause of neonatal porcine coccidiosis.20 Infected piglets develop diarrhea and
uneven weight gains.20
7.7 Treatment and Prevention
7.7.1 Treatment
In patients with AIDS and diarrhea, the use of trimethoprim–sulfamethoxazole and HAART is indicated. Treatment with HAART improves immunity, and with CD4 values >250 cells/mm3, symptoms
diminish or disappear and the excretion of oocysts is limited.83 The specific treatment is with trimethoprim (160 mg) and sulfamethoxazole (800 mg) four times per day for 10 days.5 Clinical response
with disappearance of diarrhea is observed after 48 h.69 Clinical relapses occur in 47%–50% of cases
within 8 weeks after therapy completion.69,85 Treatment of relapses is carried out with a new cycle of
trimethoprim (160 mg) and sulfamethoxazole (800 mg) and 25 mg pyrimethamine/day.29 Prophylaxis
against relapses is done with trimethoprim (160 mg) and sulfamethoxazole (800 mg) three times a week
until the CD4 cell count exceeds 250 cells/mm3.29,83 Monitoring of patients once they begin treatment
includes the number of stools per day, the daily weight, the CD4 cell count, and viral load.83 Stool examinations in 6–8 weeks or when the value of CD4 remains low or viral load increases are recommended.95
Specific treatment is indicated in AIDS patients with cystoisosporosis and acalculous cholecystitis
cystoisosporosis.6 In AIDS patients with acalculous cholecystitis, insufficient evidence is available to
determine the benefits of performing a cholecystectomy.96,97
There is no consensus on which therapy to follow for asymptomatic immunocompetent patients.
In immunocompetent patients with diarrhea and cystoisosporosis, specific treatment is performed95
with trimethoprim (160 mg) and sulfamethoxazole (800 mg) four times per day for 10 days.2 Clinical
response is observed after 48 h.98 Examination of three or more stool samples should be done after 48 h.98
Monitoring of patients with chronic diarrhea or malabsorption should include duodenal biopsy samples
for crypt hyperplasia and villous atrophy evaluation.47
The treatment of Cystoisospora infections in dogs, cats, and pigs is described in Table 7.4.2
Treatment of Cystoisosporosis in Dogs, Cats, and Pigs
Ormetoprim + sulfadimethoxine
Duration (Days)
50 mg/kg/orally
Ormetoprim (11 mg/kg) + sulfadimethoxine
(55 mg/kg)/orally
300–400 mg/kg f
20–30 mg/kg/orally
Cats, dogs
7.7.2 Prevention
Thorough washing of hands and fruits and vegetables when eating or preparing food is among the preventive measures recommended. HIV infected and those with antitumor therapy or steroids must refrain
from eating raw vegetables and fruits. They should take prophylactic trimethoprim and sulfamethoxazole treatment when necessary.
1. Levine, N.D., The Protozoan Phylum Apicomplexa, Vol. 1, Boca Raton, FL: CPR Press, 1988.
2. Lindsay, D.S., Dubey, J.P., and Blagburn, B.L., Biology of Isospora spp. from humans, nonhuman primates, and domestic animals, Clin. Microbiol. Rev., 10, 19, 1997.
3. Frenkel, J.K., Besnoitia wallacei of cats and rodents: With a reclassification of other cyst forming isosporoid coccidian, J. Parasitol., 63, 611, 1977.
4. Barta, J.R. et al., The genus Atoxoplasma (Garnham 1950) as a junior objective synonym of the genus
Isospora (Schneider 1881) species infecting birds and resurrection of Cystoisospora (Frenkel 1977) as
the correct genus for Isospora species infecting mammals, J. Parasitol., 91, 726, 2005.
5. Jongwutiwes, S., Sampatanukul, P., and Putaporntip, C. Recurrent isosporiasis over a decade in an immunocompetent host successfully treated with pyrimethamine, Scand. J. Infect. Dis., 34, 859, 2002.
6. Benator, D.A. et al., Isospora belli infection associated with acalculous cholecystitis in a patient with
AIDS, Ann. Intern. Med., 121, 663, 1994.
7. Zenta, W. and Topazian, M.D., Isospora belli cholangiopathy: Case study with histologic characterization and molecular confirmation, Hum. Pathol., 40, 1342, 2009.
8. Restrepo, C., Macher, A.M., and Radany, E.H., Disseminated extraintestinal isosporiasis in a patient with
acquired immune deficiency syndrome, Am. J. Clin. Pathol., 87, 536, 1987.
9. Comin, C.E. and Santucci, M. Submicroscopic profile of Isospora belli enteritis in a patient with acquired
immune deficiency syndrome, Ultrastruc. Pathol., 18, 473, 1994.
10. Michiels, J.F. et al., Intestinal and extraintestinal Isospora belli infection in an AIDS patient, Pathol. Res.
Pract., 190, 1089, 1994.
11. Velásquez, J.N. et al., Isosporosis and unizoite tissue cyst in patient with acquired immunodeficiency
syndrome, Hum. Pathol., 32, 500, 2001.
12. Frenkel, J.K. et al., Presença extra-intestinal de cistos unizoicos de Isospora belli em paciente com SIDA,
Rev. Soc. Bras. Med. Trop., 36, 409, 2003.
13. Dubey, J.P. and Frenkel, J.K., Extra-intestinal stages of Isospora felis and Isospora rivolta (Protozoa:
Eimeriidae) in cats, J. Protozool., 19, 89, 1972.
14. Costa, P.S. and Lopes, C.W.G., Avaliação do parasitismo por Cystoisospora felis (Wenyon, 1923)
Frenkel, 1977 (Apicomplexa: Cystoisosporinae) em coelhos do tipo carne, Rev. Bras. Parasitol. Vet.,
7, 15, 1998.
15. Melo, P. et al., Hypnozoítas de Cystoisospora felis (Wenyon, 1923) Frenkel, 1977 (Apicomplexa:
Cystoisosporinae) isolated from piglets experimentally infected, Rev. Bras. Ciência Vet., 12, 82, 2003.
16. Dubey, J.P. and Mehlhorn, H., Extraintestinal stages of Isospora ohioensis from dogs in mice,
J. Parasitol., 64, 689, 1978.
17. Massad, F.V. et al., Hypnozoítas de Cystoisospora ohioensis (Dubey, 1975) Frenkel, 1977 (Apicomplexa:
Cystoisosporinae) em frangos, Rev. Bras. Ciência Vet., 10, 57, 2003.
18. Zahed, A.A. and El-Ghaysh, A., Pig, donkey and buffalo meat as a source of some coccidian parasites
infection dogs, Vet. Parasitol., 78, 161, 1998.
19. Mitchell, S.M. et al., Cystoisospora canis Nemeséri, 1959 (syn. Isospora canis), infections in dogs:
Clinical signs, pathogenesis, and reproducible clinical disease in beagle dogs fed oocysts, J. Parasitol.,
93, 345, 2007.
20. Stuart, B.P. et al., Isospora suis enteritis in piglets, Vet. Pathol., 17, 84, 1980.
21. Stuart, B.P., Bedell, D.M., and Lindsay, D.S., Coccidiosis in swine: A search for extraintestinal stages of
Isospora suis, Vet. Rec., 110, 82, 1982.
22. Pinckney, R.D. et al., Ultrastructure of Isospora suis during excystation and attempts to demonstrate
extraintestinal stages in mice, Vet. Parasitol., 47, 225, 1993.
23. Lindsay, D.S. and Blagburn, B.L., Biology of mammalian Isospora, Parasitol. Today, 10, 214, 1994.
Biology of Foodborne Parasites
24. Jongwutiwes, S. et al., Morphologic and molecular characterization of Isospora belli oocysts from
patients in Thailand, Am. J. Trop. Med. Hyg., 77, 107, 2007.
25. Matsui, T. et al., Infectivity and sporogony of Caryospora-type oocysts of Isospora rivolta obtained by
heating, Parasitol. Res., 79, 599, 1993.
26. Roberts, W.L., Mahrt, J.L., and Hammond, D.M., The fine structure of the sporozoites of Isospora canis,
Z. Parasitenkd., 40, 183, 1972.
27. Mehlhorn, H. and Markus, M.B., Electron microscopy of stages of Isospora felis of the cat in the mesenteric lymph node of the mouse, Z. Parasitenkd., 51, 15, 1976.
28. Mitchell, S.M., Zajac, A.M., and Lindsay, D.S., Development and ultrastructure of Cystoisospora canis
Nemeséri, 1959 (syn. Isospora canis) monozoic cysts in two noncanine cell lines, J. Parasitol., 95, 793, 2009.
29. Orenstein, J.M., Isosporiasis, in: Pathology of Infectious Diseases (eds. Connor, D.H. et al.), Stamford,
CT: Appleton & Lange, 1997, pp. 1185–1190.
30. Scholtyseck, E. and Mehlhorn, H., Fine structure of macrogametes and oocysts of coccidia and related
organisms, Z. Parasitenk., 37, 1, 1971.
31. Dubey, J.P., Lindsay, D.S., and Lappin, M.R., Toxoplasmosis and other intestinal coccidial infections in
cats and dogs, Vet. Clin. Small Anim., 39, 1009, 2009.
32. Votýpka, J. et al., Molecular phylogenetic relatedness of Frenkelia spp. (Protozoa, Apicomplexa) to
Sarcocystis falcatula Stiles, 1893: Is the genus Sarcocystis paraphyletic? J. Eukaryot. Microbiol., 45,
137, 1998.
33. Barta, J.R. et al., Phylogenetic relationships among eight Eimeria species infecting domestic fowl
inferred using complete small subunit ribosomal DNA sequences, J. Parasitol., 83, 262, 1997.
34. Carreno, R.A. and Barta, J.R., An Eimeriid origin of isosporoid coccidia with Stieda bodies as shown by
phylogenetic analysis of small subunit ribosomal RNA gene sequences, J. Parasitol., 85, 77, 1999.
35. Samarasinghe, B., Johnson, J., and Ryan, U., Phylogenetic analysis of Cystoisospora species at the rRNA
ITS1 locus and development of a PCR-RFLP assay, Exp. Parasitol., 118, 592, 2008.
36. Dubey, J.P., Life-cycle of Isospora ohioensis in dogs, Parasitology, 77, 1, 1978.
37. Dubey, J.P., Life-cycle of Isospora rivolta (Grassi, 1879) in cats and mice, J. Protozool., 126, 433, 1979.
38. Ash, L.R. and Orihel, T.C., Parasites: A Guide to Laboratory Procedures and Identification, Chicago, IL:
American Society of Clinical Pathologist, 1991.
39. Brandborg, L.L., Goldberg, S.B., and Breidenbach, W.C., Human coccidiosis: A possible cause of malabsorption, N. Engl. J. Med., 283, 1306, 1970.
40. Faust, E.C. et al., Human isosporosis in the Western hemisphere, Am. J. Trop. Med. Hyg., 10, 343, 1961.
41. Ma, P., Kaufman, D., and Montana, J., Isospora belli diarrheal infection in homosexual men, AIDS Res.,
1, 327, 1984.
42. Miller, F.H., Pizzuto, A.V., and McCauley, H., Human isosporosis: Two cases, Am. J. Trop. Med. Hyg.,
20, 23, 1971.
43. Soave, R. and Johnson, W.D., Cryptosporidium and Isospora belli infections, J. Infect. Dis., 157, 225, 1988.
44. Varela, M. et al., Fuchsin fluorescence and autofluorescence in Cryptosporidium, Isospora and Cyclospora
oocysts, Int. J. Parasitol., 28, 1881, 1998.
45. Ng, E. et al., Demonstration of Isospora belli by acid-fast stain in a patient with acquired immune deficiency syndrome, J. Clin. Microbiol., 20, 384, 1984.
46. Dammin, G.J. and Dooley, J.R., Coccidiosis, in: Pathology of Tropical and Extraordinary Diseases
(eds. Chapman, H. and Connor, D.H.), Vol. 1, Section 7, Protozoa, Chapter 13, Washington, DC: Armed
Forces Institute of Pathology, 1976, pp. 332–335.
47. Trier, J.S. et al., Chronic intestinal coccidiosis in man: Intestinal morphology and response to treatment,
Gastroenterology, 66, 923, 1974.
48. Liebman, W.M. et al., Intractable diarrhea of infancy due to intestinal coccidiosis, Gastroenterology, 78,
579, 1980.
49. Warnnissorn, N. et al., Isospora belli infection associated with chronic cholecystitis and sclerosing cholangitis in immunocompetent host: A case report, Siriraj Hosp. Gaz., 55, 419, 2003.
50. Walther, Z. and Topazian, M.D., Isospora cholangiopathy: Case study with histologic characterization
and molecular confirmation, Hum. Pathol., 40, 1342, 2009.
51. Müller, A. et al., Detection of Isospora belli by polymerase chain reaction using primers based on smallsubunit ribosomal RNA sequences, Eur. J. Clin. Microbiol. Infect. Dis., 19, 631, 2000.
52. ten Hove, R.J. et al., Real-time polymerase chain reaction for detection of Isospora belli in stool samples,
Diagn. Microbiol. Infect. Dis., 61, 280, 2008.
53. Taniuchi, M. et al., Multiplex PCR method to detect Cyclospora, Cystoisospora, and Microsporidia in
stool samples, Diagn. Microbiol. Infect. Dis., 71, 386, 2011.
54. Murphy, S.C. et al., Molecular diagnosis of cystoisosporiasis using extended-range PCR screening,
J. Mol. Diagn., 13, 359, 2011.
55. Carreno, R.A. et al., Phylogenetic analysis of coccidia based on 18S rDNA sequence comparison indicates
that Isospora is most closely related to Toxoplasma and Neospora, J. Eukaryot. Microbiol., 45, 184, 1998.
56. Franzen, C. et al., Taxonomic position of the human intestinal protozoan parasite Isospora belli as based
on ribosomal RNA sequences, Parasitol. Res., 86, 669, 2000.
57. Millar, B.C. et al., Sequence analysis of partial regions of the 5.8S rRNA internal transcribed region 2 and
28S rRNA of Isospora belli, Br. J. Biomed. Sci., 60, 114, 2003.
58. Velásquez, J.N. et al., Molecular characterization of Cystoisospora belli and unizoite tissue cyst in
patients with acquired immunodeficiency syndrome, Parasitology, 138, 279, 2011.
59. Resende, D.V. et al., Polymorphisms in the 18S rDNA gene of Cystoisospora belli and clinical features
of cystoisosporosis in HIV-infected patients, Parasitol. Res., 108, 679, 2011.
60. Ruttkowski, B., Joachim, A., and Daugschies, A., PCR-based differentiation of three porcine Eimeria
species and Isospora suis, Vet. Parasitol., 95, 17, 2001.
61. Johnson, J. et al., Molecular identification and prevalence of Isospora sp. in pigs in Western Australia
using a PCR-RFLP assay, Exp. Parasitol., 120, 191, 2008.
62. Lalonde, L.F. and Gajadhar, A.A., Detection and differentiation of coccidian oocysts by real-time PCR
and melting curve analysis, J. Parasitol., 97, 725, 2011.
63. Matsubayashi, M. et al., Phylogenetic identification of Cystoisospora spp. from dogs, cats, and raccoon
dogs in Japan, Vet. Parasitol., 176, 270, 2011.
64. He, P. et al., Cystoisospora spp. from dogs in China and phylogenetic analysis of its 18S and ITS1 gene,
Vet. Parasitol., 190, 254, 2011.
65. Sagua, H. et al., Algunos aspectos clínicos y epidemiológicos de la isosporosis intestinal. Estudio en 26
pacientes pediátricos de la ciudad de Antofagasta, Rev. Chil. Pediatr., 50, 15, 1979.
66. Sorvillo, F.J. et al., Epidemiology of isosporiasis among persons with acquired immunodeficiency syndrome in Los Angeles County, Am. J. Trop. Med. Hyg., 53, 656, 1995.
67. de Oliveira-Silva, M.B. et al., Seasonal profile and level of CD4+ lymphocytes in the occurrence of cryptosporidiosis and cystoisosporidiosis in HIV/AIDS patients in the Triângulo Mineiro region, Brazil, Rev.
Soc. Bras. Med. Trop., 40, 512, 2007.
68. Gupta, S. et al., Chronic diarrhoea in HIV patients: Prevalence of coccidian parasites, Indian J. Med.
Microbiol., 26, 172, 2008.
69. DeHovitz, J.A. et al., Clinical manifestations and therapy of Isospora belli infection in patients with the
acquired immunodeficiency syndrome, N. Engl. J. Med., 315, 87, 1986.
70. Kirkpatrick, C.E. and Dubey, J.P., Enteric coccidial infections with Isospora, Sarcocystis, Cryptosporidium,
Besnoitia and 1987, Vet. Clin. North Am. Small Anim. Pract., 17, 1405, 1987.
71. Niestrath, M. et al., The role of Isospora suis as a pathogen in conventional piglet production in Germany,
J. Vet. Med. B Infect. Dis. Vet. Public Health, 49, 176, 2002.
72. Palmer, C.S. et al., National study of the gastrointestinal parasites of dogs and cats in Australia, Vet.
Parasitol., 151, 181, 2008.
73. Barutzki, D. and Schaper, R., Results of parasitological examinations of faecal samples from cats and
dogs in Germany between 2003 and 2010, Parasitol. Res., 109, S45, 2011.
74. da Cunha Ferreira, R. et al., Changes in the rate of crypt epithelial cell proliferation and mucosal morphology induced by a T-cell-mediated response in human small intestine, Gastroenterology, 98, 1255,
75. Hand, T.W. et al., Acute gastrointestinal infection induces long-lived microbiota-specific T cell responses,
Science, 337, 1553, 2012.
76. Worliczek, H.L. et al., Changes in lymphocyte populations in suckling piglets during primary infections
with Isospora suis, Parasite Immunol., 32, 232, 2010.
77. Cornelissen, J.B. et al., Host response to simultaneous infections with Eimeria acervulina, maxima and
tenella: A cumulation of single responses, Vet. Parasitol., 162, 58, 2009.
Biology of Foodborne Parasites
78. Sühwold, A. et al., T cell reactions of Eimeria bovis primary and challenge-infected calves, Parasitol.
Res., 106, 595, 2010.
79. Pogonka, T. et al., CD8+ cells protect mice against reinfection with the intestinal parasite Eimeria falciformis, Microbes Infect., 12, 218, 2010.
80. Aleksandersen, M. et al., Scarcity of gamma delta T cells in intestinal epithelia containing coccidia
despite general increase of epithelial lymphocytes, Vet. Pathol., 32, 504, 1995.
81. Brenchley, J.M. and Douek, D.C., HIV infection and the gastrointestinal immune system, Mucosal
Immunol., 1, 23, 2008.
82. Ryan, E.T., Cronin, C.G., and Branda, J.A., Case 38–2011. A 34-year-old man with diarrhea and weakness, N. Engl. J. Med., 365, 2306, 2011.
83. Kartalija, M. and Sande, M., Diarrea and AIDS in the era of highly active antiretroviral therapy, Clin.
Infect. Dis., 28, 701, 1999.
84. Bollinger, R. and Quinn, T., Approach to gastrointestinal infection in immunosuppressed, in:
Gastrointestinal and Hepatic Infections (eds. Surawicz, C. and Owen, R.), St. Louis, MO: W.B. Saunders
Company, 1995, pp. 575–596.
85. Pape, J.W., Verdier, R.I., and Johnson, W.D., Treatment and prophylaxis of Isospora belli infection in
patients with the acquired immunodeficiency syndrome, N. Engl. J. Med., 320, 1044, 1989.
86. Henderson, H.E. et al., The human Isospora, Am. J. Hyg., 78, 302, 1963.
87. Connal, A., Observations on the pathogenicity of Isospora hominis, Rivolta, emend. Dobell, based on a
second case of human coccidiosis in Nigeria, with remarks on the significance of Charcot-Leyden crystals in the faeces, Trans. Roy. Soc. Trop. Med. Hyg., 16, 233, 1922.
88. Jarpa, G.A., Coccidiosis humana, Biologica (Santiago), 39, 33, 1966.
89. Barksdale, W. and Routh, C.F., Isospora hominis infection among American personnel in the Southwest
Pacific, Am. J. Trop. Med., 28, 639, 1948.
90. Resiere, D. et al., Isospora belli infection in a patient with non-Hodgkin’s lymphoma, Clin. Microbiol.
Infect., 9, 1065, 2003.
91. Greenberg, S.J. et al., Isospora belli enteric infection in patients with human Tcell leukemia virus type
I-associated adult T-cell leukemia, Am. J. Med., 85, 435, 1988.
92. Malik, S. et al., Refractory isosporiasis, Indian J. Pediatr., 72, 437, 2005.
93. Koru, O. et al., Isospora belli infection in a renal transplant recipient, Turkiye Parazitol. Derg., 31, 98,
94. Atambay, M. et al., A rare diarrheic parasite in a liver transplant patient: Isospora belli, Transplant. Proc.,
39, 1693, 2007.
95. Carr, A. et al., Treatment of HIV-1-associated microsporidiosis and cryptosporidiosis with combination
antiretroviral therapy, Lancet, 351, 256, 1998.
96. Knapp, P.E., Saltzman, J.R., and Fairchild, P., Acalculous cholecystitis associated with microsporidial
infection in a patient with AIDS, Clin. Infect. Dis., 22, 195, 1996.
97. French, A.L. et al., Cholecystectomia in patients with AIDS: Clinicopathologic correlations in 107 cases,
Clin. Infect. Med., 21, 852, 1995.
98. Westerman, E.L. and Christensen, R.P., Chronic Isospora belli infection treated with co-trimoxazole,
Ann. Int. Med., 91, 413, 1979.
Sandipan Ganguly
Morphology and Life Cycle...........................................................................................................132
8.2.1 Morphology.......................................................................................................................132
8.2.2 Life Cycle..........................................................................................................................133
8.3 Biology, Genetics, and Genomics................................................................................................. 134
8.3.1 Biochemistry and Metabolism..........................................................................................135 Carbohydrate Metabolism.................................................................................135 Electron Transport.............................................................................................135 Lipid Metabolism.............................................................................................. 136 Protein Synthesis.............................................................................................. 136 Nucleic Acid Metabolism.................................................................................. 136
8.3.2 Genetics and Genomics.................................................................................................... 136
8.4 Diagnosis and Typing.................................................................................................................... 137
8.4.1 Diagnosis.......................................................................................................................... 137
8.4.2 Coding Loci as Genotyping Markers............................................................................... 138
8.4.3 Noncoding Loci as Genotyping Markers......................................................................... 138
8.5 Epidemiology and Molecular Epidemiology................................................................................ 139
8.5.1 Foodborne and Waterborne Outbreaks of Entamoeba.................................................... 139
8.5.2 Molecular Epidemiology.................................................................................................. 140 Genotyping Based on Conventional Polymorphic Markers............................. 140 tRNA-Linked STR Loci and Its Association with Virulence.......................... 140 Nonrepetitive Coding Loci Significantly Linked with Parasite Virulence.......141 Transposable Elements and Genomic Rearrangements.....................................141 Gene Families and Diversity.............................................................................141 Multiple Infections and the Need for Sequencing.............................................142
8.6 Pathogenesis and Clinical Features...............................................................................................142
8.6.1 Clinical Features...............................................................................................................142 Symptomatic Infection.......................................................................................142 Intestinal Infections...........................................................................................142 Extraintestinal Infection....................................................................................143 Asymptomatic Infection....................................................................................143
8.7 Treatment and Prevention..............................................................................................................143
8.7.1 Treatment...........................................................................................................................143
8.7.2 Prevention......................................................................................................................... 144
References............................................................................................................................................... 144
Biology of Foodborne Parasites
8.1 Introduction
Amoebiasis, an invasive intestinal or extraintestinal infection caused by the protozoan parasite
Entamoeba histolytica, is responsible for approximately 100,000 deaths annually. It is regarded as
the fourth leading cause of death due to a protozoan infection after malaria, Chagas’ disease, and
leishmaniasis and the third cause of morbidity in this organism group after malaria and trichomoniasis.1 The earliest mention of amoebiasis as a bloody, mucus diarrhea was possibly found in the
Sanskrit document Bhrigu Samhita written about 1000 BC.2 As amoebiasis became widespread in
the developed world, there were numerous records of “bloody flux” in Europe, Asia, Persia, and
Greece in the Middle Ages.3 The disease appears to have been introduced into the New World by
Europeans sometime in the sixteenth century.4 James Annersley (1828) first recognized the relationship between dysentery and liver abscess, which was thoroughly described later by William Budd
(1857). Losch discovered amoeba in 1875 and also established the relationship between the parasite
and the disease.2 Schaudinn (1903) named E. histolytica, and Craig (1905) confirmed the findings of
Schaudinn, and finally the pathogenicity of these amoebae was proven by the experiments conducted
by Walker and Sellards (1913). Entamoeba dispar, an amoeba morphologically similar to E. histolytica, which also colonizes the human gut with no invasive potential, was recognized as a separate
species in 1925.5 Together, E. histolytica and E. dispar infect about 10% of the world’s p­ opulation.1
Another species, Entamoeba moshkovskii, also has considerable prevalence in E. ­histolytica endemic
8.2 Morphology and Life Cycle
8.2.1 Morphology
The life cycle of E. histolytica includes several stages: trophozoite, precyst, cyst, and metacyst.6
Trophozoites found in tissues vary in size from 12 to 60 µm in diameter, while those found in
nondysenteric stools or in cultures measure 7–30 µm. The ectoplasm is clear, and the endoplasm
is granular and may contain bacteria and/or erythrocytes in various stages of digestion. The trophozoite nucleus is spherical and varies from 4 to 7 µm. In stained preparation, the nucleus shows
(1) a nuclear membrane, which is delicate and lined with a single layer of uniformly distributed
fine chromatin granules; (2) a karyosome, which is small and dot like, central in position, and surrounded by a clear halo; and (3) the space between the karyosome and the nuclear membrane, which
is traversed by a fine thread of “linin” network having a spoke-like radial arrangement. The organism does not contain a cytochrome system or a distinct rough endoplasmic reticulum and Golgi
apparatus. However, immunofluorescence analysis using antibodies to proteins located in the rough
endoplasmic reticulum and Golgi apparatus in other eukaryotes has identified a putative endoplasmic reticulum and Golgi structures in E. histolytica.7 Although Entamoeba does not apparently
have a structure that resembles mitochondria, immunolocalization has revealed that a functional
mitochondrial remnant called a “crypton”8 or “mitosome” 9 is present in the cytoplasm. Under the
microscope, mitosomes are ovoid structures smaller than 0.5 µm in diameter. Since the number of
identifiable mitosomal proteins is very small, a comprehensive insight into the organelle’s function
is still unknown. Genes encoding mitochondrial-type chaperonins (cpn60, hsp10, and mt-hsp70)
have been identified and appear to be synthesized with amino-terminal signal sequences.10 The
important machinery has been shown to be conserved with that in true mitochondria, but none of
the proteins involved in mitosomal protein have been identified with certainty. Cytoskeleton components such as actin,11 tubulin,12 cytoplasmic microfilaments, nuclear microtubules, myosin,13 and
a β1 integrin-like molecule14 have been described. Ribosomes form aggregated crystalline arrays in
the cytoplasm of the trophozoite.
8.2.2 Life Cycle
Cysts and trophozoites are passed in feces (Figure 8.1). Cysts are the infective stage of the life cycle and
are mainly found in the stool, especially in formed stools, whereas trophozoites are typically absent, but
may be present in diarrheal stool. Infection with E. histolytica occurs by ingestion of mature cysts in fecal-­
contaminated water or food and also through contaminated hands. The cysts then excyst in the small intestine
in acidic pH, releasing trophozoites—the pathogenic stage of the life cycle of Entamoeba. The trophozoites
can migrate to the large intestine and, in this basic pH environment, multiply by binary fission and produce
cysts, which are passed in the feces (Figure 8.2). Mature cysts can survive for weeks in the environment.
The trophozoites can remain confined in the intestinal lumen without causing any disease outcome, passing cysts in the stool. However, in some cases, the trophozoite invades the intestinal mucosa and enters the
bloodstream and progresses to extraintestinal sites such as liver, brain, and lungs resulting in clinical manifestations. Transmission can also occur through exposure to fecal matter during sexual contact.15
In the precyst stage, the trophozoite becomes approximately the same size as the cyst. The cytoplasm
is cleared of all food inclusions but usually contains diffuse glycogen deposits and occasional chromatoid bodies, made up of ribosomes. The precystic form is uninucleate, and the enlarged nucleus contains
a karyosome that is more or less eccentric. Cysts are round or slightly oval, ranging between 8 and
14 µm. The cyst wall contains chitin as shown by chemical analysis and x-ray diffraction studies of purified preparations.16 One to four nuclei are clearly seen, and the nuclear membrane is uniformly lined with
peripheral chromatin. The karyosome is small, usually centrally located within the nucleus. Generally,
glycogen and chromatoidal materials disappear as the cyst matures. During the process of excystation,
the encysted amoeba becomes very active, separating from the cyst wall. The quadrinucleate amoeba
escapes from the cyst wall through a tiny pore, and the nuclei clump together.17
Invasion/colonization sites
Mature cyst
Immature cyst
Human host
FIGURE 8.1 Life cycle of E. histolytica. Cysts in stools initiate infection in a healthy person. The quadrinucleate cysts
enter the host through oral route. The mature cysts with four nuclei then undergo excystation within the host body to release
trophozoites. The trophozoites then colonize in the intestine and may sometime migrate to extraintestinal organs like the
lung and liver. They multiply in number through binary fission. Trophozoites can also undergo encystation, and the cysts
are passed through the host with the stool, initiating new infections.
Biology of Foodborne Parasites
FIGURE 8.2 Morphology. Structure of trophozoite and cyst of (a and b) E. dispar and (c and d) E. histolytica. Both species
have the morphologically identical structure in wet mount microscopy.
8.3 Biology, Genetics, and Genomics
Several species of Entamoeba are found in humans. There are harmless species such as E. coli and
E. dispar, while E. histolytica is the pathogen responsible for amoebic dysentery and amoebic liver
abscesses (ALAs). All the Entamoeba species are found in the intestines of the animals they infect,
the only exception being E. gingivalis, which lives in the mouth, and E. moshkovskii, which for a long
time was considered as a free-living amoeba; but in the last decade, it has been demonstrated that
E. moshkovskii can infect humans and can be found more frequently in regions where amoebiasis
is endemic, becoming a challenge to differentiate it from the E. histolytica/E. dispar complex.18 E.
invadens is a species that is specific for reptiles. In contrast to other species, E. invadens can form
cysts in vitro.19
E. histolytica trophozoites live and multiply indefinitely within the mucosa of the large intestine
feeding normally on starches and mucous secretions and interacting metabolically with gut bacteria.
However, such trophozoites commonly initiate tissue invasion when they hydrolyze mucosal cells and
absorb the predigested products in order to meet their dietary provisions. Once the parasites invade the
intestinal wall, they reach the submucosa and the underlying blood vessels. From there, trophozoites
travel in the blood to extraintestinal sites such as the liver, lungs, skin, and even brain.
The signals leading to encystation or excystation are poorly understood, but findings from E. invadens
suggest that ligation of a surface galactose-binding lectin on the surface of the parasite might be the
trigger for encystation.20 Also, several previous proteomic and transcriptomic studies have shown that
a few dozen Rab genes/proteins are involved in important biological processes, such as stress response,
virulence, pathogenesis, and stage conversion.21
8.3.1 Biochemistry and Metabolism
E. histolytica is one of the primitive eukaryotes; thus, its biochemistry has a number of unusual aspects
that are evocative of prokaryotes. Glycolysis in E. histolytica lacks conventional eukaryotic glycolytic
enzymes22 and utilizes PPi instead of ATP in a number of enzyme reactions, for example, phosphofructokinase. Another unusual feature is the absence of pentose phosphate shunt. Amoebae resemble more
closely anaerobic and microaerophilic bacteria. They are able to consume oxygen despite the lack of
mitochondria and can grow in an atmosphere containing up to 5% oxygen. Below this condition, amoebae are able to detoxify the products of oxygen reduction. Carbohydrate Metabolism
Glucose and, to a lesser extent, galactose are the main sources of energy for E. histolytica. Glucose
uptake occurs via a specific transport system that provides approximately 100 times the amount incorporated by endocytosis.23 This transport system is the rate-limiting step in glucose consumption. Although
no free glucose has been found in E. histolytica, glucokinase activity is 20 times greater than the rate of
glucose uptake. Glycogen is the main form of glucose storage, and three enzymes are involved in glycogen metabolism, namely, UDP glucose pyrophosphorylase, phosphoglucomutase, and phosphorylase.
Phosphorylase is involved in glycogen synthesis.24
Amoebic catabolism of glucose differs considerably from that of most eukaryotic cells, given the presence of uncommon glycolytic enzymes and the absence of mitochondria, cytochromes, and a citric acid
cycle. The parasite degrades glucose to pyruvate via a modified Embden–Meyerhof pathway.25 Lactate
is not an end product, and lactate dehydrogenase has not been reported. Pyruvate is converted mostly to
ethanol, even in the presence of oxygen, via coenzyme A and pyruvate oxidase.26 Glucose 6-phosphate is
either produced directly from the phosphorylation of glucose by glucokinase or comes from stored glycogen via glucose 1-phosphate and phosphoglucomutase. In the presence of phosphoglucoisomerase, glucose 6-phosphate produces fructose 6-phosphate, which subsequently forms fructose 1,6-diphosphate.
Rather than using the usual ATP-dependent phosphofructokinase, Entamoeba is found to possess an
inorganic pyrophosphate (PPi)-dependent enzyme catalyzing the following reaction:
Fruct-6-PO4 + PPi ⇌ Fruct-1,6-diPO4 + Pi
Fructose 1,6-diphosphate forms phosphoenolpyruvate via the usual glycolytic sequences. Due to the
absence of the pyruvate kinase, the conversion of phosphoenolpyruvate to pyruvate is catalyzed by the
PPi-dependent enzyme pyruvate phosphate dikinase:
Pyruvate + ATP + Pi ⇌ Phosphoenolpyruvate + AMP + PPi
It should be noted that glucose 6-phosphate dehydrogenase has not been found in E. histolytica and is
not degraded by the pentose phosphate pathway.27 Electron Transport
Proteins with heme groups like cytochrome are absent in E. histolytica, yet nonheme iron and acid labile
sulfur have been reported in trophozoites.28 During aerobiosis, electrons are transferred from reduced
substrates via a succession of carriers, including flavins, ferredoxin, other FeS proteins, and ubiquinone,
to molecular oxygen, which is reduced to water. Since elevation of oxygen tension is toxic to this parasite,
high concentrations of cysteine and other thiols play an important role in the antioxidative defenses by
the parasite. An iron-dependent superoxide dismutase, an oxidative stress-detoxifying enzyme, is also
present in E. histolytica.29
Biology of Foodborne Parasites Lipid Metabolism
Parasites in general possess only limited ability to synthesize long-chain fatty acids de novo and, therefore, rely to a great extent on the host to provide for their essential lipid development. The most abundant
sterol in Entamoeba is cholesterol, which the parasite acquires from the host. Cholesterol is enriched
with phospholipids in the plasma membrane. Phosphatidylethanolamine predominates over phosphatidylcholine in the plasma membrane, although the latter is the most abundant lipid in internal vesicles. An
unusual phospholipid, ceramide aminoethyl phosphonate, has been demonstrated in internal vesicles but
is mostly concentrated in the plasma membrane. The unusual composition of the membrane lipids might
account, at least in part, for the pronounced plasticity and stability of the membranes. Furthermore, this
may have biological importance, as this compound is resistant to hydrolysis and may confer protection to hydrolytic enzymes present in the host’s gastrointestinal tract, as well as to the action of its own
phospholipase.26 Protein Synthesis
Sulfur-containing amino acid metabolism in E. histolytica is unique in a variety of aspects including a
lack of both forward and reverse transsulfuration pathways; a lack of enzymes responsible for cysteine
and homocysteine degradation in mammals, including cysteine dioxygenase and phosphopantothenoylcysteine synthetase; and the presence of de novo sulfur-assimilatory cysteine biosynthetic pathway.30
In E. histolytica, cysteine biosynthesis is coordinately regulated with serine metabolism: serine is a
precursor for cysteine synthesis. Nucleic Acid Metabolism
Presently, very limited information is available on nucleic acid metabolism in Entamoeba. There is a
de novo synthesis of purine bases, and pyrimidine synthesis is based on orotic acid incorporation. Uptake
of both purine and pyrimidine bases and nucleosides is, in part, carrier mediated. There are two purine
uptake sites, one for adenine and adenosine and the other for adenosine and guanine. Passive uptake of
guanine, inosine, and hypoxanthine is also observed. Two pyrimidine uptake sites are observed, one for
uridine/cytidine and the other for uridine and a purine nucleoside, adenosine.26
8.3.2 Genetics and Genomics
The haploid genome of E. histolytica strain HK-9 is 3 × 107 bp31 in size based on renaturation kinetics
study, and the ploidy has been calculated between 8 and 10.23 The exact chromosome number of E. histolytica
is difficult to determine because of its ability to change ploidy number and due to the high content of
repetitive DNA in its genome. Expansion and contraction of subtelomeric repeats are responsible for this
varying chromosome size. Interestingly, these subtelomeric regions consist of tRNA-containing arrays.
The amoeba genome is low in G+C content (about 22.4%). The ploidy appears to be variable within a cell
lineage under different growth conditions and life cycle phases.32 Molecular karyotyping study revealed
variations of chromosome size among different strains of E. histolytica as well as the presence of multiple circular DNA structure along with the linear DNA.33 The rRNA genes are present in one of such
circular DNA molecules that exist in multiple copies per nucleus. This circular DNA molecule might
be important for determining the parasite phenotype. The 9938 genes with an average size of 1.17 kb
constitute 49% of the total genome. One-fourth of the genes are predicted to have introns, and about 6%
of genes possess multiple introns.34
In 2005, a draft of the complete genome of E. histolytica was published34 making it one of the first
protist genomes to be sequenced. The genome project was initiated in 2000 at the Wellcome Trust Sanger
Institute and The Institute for Genomic Research (TIGR) in the United Kingdom and the United States,
respectively. The presence of the multiple copies of an episome encoding the rRNA genes has complicated the sequencing process because ~15% of the resulting sequence corresponds to this episome.
A library, relatively free of rDNA circles, has been derived at the Sanger Centre by linearizing the rDNA
circles with a restriction digestion by PpoI and then by physically removing the linearized DNA from the
pulse-field gel. This library was compared with the sequences obtained at TIGR to ensure that nothing
apart from rDNA circles has been lost.
Initial characterization of the sequences shows that the genome is highly repetitive, with 40% of the
sequence assigned to repetitive elements. Thirteen percent of the 49,000 sequences examined contain
tRNA genes, most of which are located in tandem arrays.35 The predominant location of tRNA arrays at
the telomere could account for the variation in sizes of the homologous chromosomes. Another finding
has been the identification of a gene family of >100 proteins containing the CXXC motif, which includes
the intermediate subunit of the E. histolytica Gal/GalNAc lectin.36
Recently, all sequence assemblies over 10 kb in length from a 7X assembly of the Entamoeba genome
have been annotated. These assemblies comprise of approximately 12 Mb of nonrepetitive sequence data
and a majority of the coding regions of E. histolytica.
An unusual feature of genomic organization of E. histolytica is the presence of 4500 copies of tandemly arrayed tRNA genes, which comprise of short tandemly repeated (STR) sequences in their intergenic region.37 Interspecific and interstrain variations of the size, number, and orientation of these STRs
are observed in Entamoeba. Genome rearrangement might be one of the possible reasons for these STR
variations, which can also lead to the differential disease outcome by the same Entamoeba species.38
Abundance of transposons and repetitive DNA can also play an important role in the genomic rearrangement of Entamoeba. A comprehensive study of three Entamoeba genomes leads to the identification of
hundreds of copies of long interspersed elements (LINEs) and short interspersed elements (SINEs) along
with Entamoeba-specific repeats. LINEs are class I transposons having their own reverse transcriptase
and other activities essential for retrotransposition, whereas SINEs are class II transposons that lack
the essential machinery to replicate themselves and are reverse transcribed by a reverse transcriptase
encoded by LINE. The LINEs and SINEs are usually more prevalent in E. invadens and E. moshkovskii
and are rare in E. histolytica and E. dispar.39
E. histolytica also contains large multigene families; among them, important are AIG1-like GTPases
and Rab GTPases. Although the function of AIG1-GTPases is precisely unknown, differential expression suggests that they may be associated with virulence and adaptation. In contrast, Rab GTPases have
a conserved machinery for controlling the vesicular trafficking.39
8.4 Diagnosis and Typing
8.4.1 Diagnosis
Diagnosis of E. histolytica has historically relied on microscopic examination. However, lack of trained
microscopists, delayed delivery of specimens, and difficulties in differentiation of nonmotile trophozoites from polymorphonuclear leukocytes, macrophages, and other tissue cells are some of the limitations
of microscopy.40
Visual examination of the colon using colonoscopy remains a very useful diagnostic method to eliminate noninfectious causes of bloody diarrhea and to confirm uncertain diagnoses of amoebic colitis.
When colonic biopsies are stained with periodic acid-Schiff, amoebic trophozoites appear magenta in
color and are clearly visible at the base of ulcerations.
Immunological methods including enzyme-linked immunosorbent assay (ELISA), enzyme immunoassay (EIA), indirect hemagglutinin assay (IHA), indirect immunofluorescent antibody (IFA) (Figure 8.3),
latex agglutination (LA), agar gel diffusion (AGD), and counterimmunoelectrophoresis (CIE) are also
used for diagnosis.41 Stool antigen detection tests are more specific than microscopy, as they distinguish
E. histolytica from E. dispar. Several antigen-based ELISA kits are now commercially available to distinguish these two species. They are sensitive, specific, and easy to perform. However, all these tests have
different limitations such as the inability to use fixed or frozen specimens. Detection of circulating antigen
in the serum is a promising yet still experimental approach to the diagnosis of amoebic liver abscess.40
Several PCR-based methods that amplify E. histolytica DNA in stool samples have also been developed.17,26 The specificity and sensitivity are high, and diagnosis is very accurate. A real-time PCR
Biology of Foodborne Parasites
FIGURE 8.3 Immunofluorescence microscopy. Immunofluorescence microscopy of E. histolytica cyst using an antibody
against a cell surface protein.
assay was also developed for sensitive and specific detection and differentiation of E. histolytica and
E. dispar in human feces.42 Given the high sensitivity and specificity of the developed PCR assay and
the inability of microscopy to distinguish between the two amoeba species, PCR-based methods are
more suitable than microscopy or immunological methods to correctly diagnose intestinal E. histolytica or E. dispar infection.
However, PCR-based detection methods primarily exploit the genetic differences between these two
morphologically identical Entamoeba species. E. histolytica is significantly different from its morphologically identical sibling E. dispar in both coding and noncoding regions. Among the polymorphic
coding regions, serine-rich E. histolytica protein (SREHP) and chitinase are particularly important. The
noncoding regions like tRNA-linked STR are also found to be useful.43 Since they are linked with species evolution, they have been used as genetic markers for interspecific differentiation.
8.4.2 Coding Loci as Genotyping Markers
SREHP gene and chitinase gene are the two most widely used genetic markers. SREHP is an immunodominant surface antigen containing tandem repeats of related dodeca- and octapeptides.44 Sequence
analysis of PCR-amplified target gene45,46 indicates definite differences in repeat types, numbers, and
arrangement patterns among two closely related Entamoeba species. However, noncoding tRNA-linked
STR loci have been preferred over conventional genetic markers for their increasing evolutionary
Chitinase is also a repeat-containing gene expressed only during encystation of the parasite.48 Although
this gene is present in both E. histolytica and E. dispar, genetic organization differs between the species.
Other genetic markers such as the Gal/GalNAc lectin have also been used.49
8.4.3 Noncoding Loci as Genotyping Markers
Noncoding tRNA-linked STR loci have been preferred for their evolutionary significance and distinct interspecific structural variations.50 E. histolytica has two versions of NK arrays, (NK1) and
(NK2), while E. dispar contains only one type. Thus, E. dispar has 24 arrays rather than 25 identified in E. ­histolytica. Moreover, E. moshkovskii arrays are significantly smaller than their homolog in
E. ­histolytica and E. dispar and do not contain any STRs in their intergenic regions.35
8.5 Epidemiology and Molecular Epidemiology
Amoebiasis occurs worldwide and especially in tropics. Its distribution in developed and developing
countries is mainly a reflection of cultural habits, socioeconomic status, and level of sanitation and
overcrowding. It has remained an important clinical problem with a significant mortality and one of the
major causes of traveller’s diarrhea. About one-tenth of the world population is stated to be at risk of
infection with E. histolytica, resulting in up to 100,000 deaths worldwide each year.51 The majority of
deaths were a consequence of severe complications associated with intestinal or extraintestinal invasive
disease. Recent reports show that E. moshkovskii is also common in some areas of E. histolytica endemicity.52 Among children, the mortality rate is less than 1% for those with mild amoebic dysentery and
40% when gastrointestinal perforation or peritonitis is present.50 The disease is most prevalent in tropical and subtropical countries where colonization with E. histolytica has been observed in 5% or more of
poor children. Less than 10% of colonized individuals develop colitis, with the rest clearing the infection
within months.51 Immigration and travel between countries may increase the incidence of disease worldwide. Countries with the highest E. histolytica endemicity include Mexico, India, Eastern and Southern
Africa, and parts of Central and South America. In Mexico, one in five infected patients develops invasive amoebiasis.51 The prevalence of invasive amoebiasis is equal among males and females; however,
extraintestinal dissemination occurs 3–10 times more often in the male population. The prevalence of
amoebiasis in the homosexual male population is reported to be between 20% and 30%. E. dispar is
the predominant species infecting this group.15 Infection with the commensal E. dispar is much more
common. Studies from Bangladesh44 have shown that even in endemic areas, E. dispar is more common
than E. histolytica. However, decreasing occurrence of Entamoeba infection has also been reported from
tropical endemic areas like India.53 Apart from these three species, several other species of Entamoeba
are present in humans, such as Entamoeba coli, E. hartmanni, E. gingivalis, E. polecki, and E. chattoni.
Some of them also infect animals.54
8.5.1 Foodborne and Waterborne Outbreaks of Entamoeba
Amoebiasis is one of the major water- and foodborne diseases, affecting about 12% of the world’s population, and is also a major cause of traveller’s diarrhea. Natural contamination of drinking water with sewage
leads to the easy transport of Entamoeba cysts. Inferior hygiene condition and inadequate knowledge of
water treatment in general population, especially in the developing countries, lead to the outbreak of amoeba
infection. The prevalence of infection varies between 1% in industrialized countries to between 50% and
80% in tropical countries, where transmission of E. histolytica by untreated drinking water is common.
One of the most severe outbreaks is the World’s Fair outbreak in Chicago in 1933 due to drinking
contaminated water, sickening about 1000 person and killing 58 of them.55 More recently, an outbreak
has occurred in Republic of Georgia in 1998 with 177 reported cases, including 106 liver abscess cases.56
Amoebiasis outbreaks in jails, institutions, day-care centers, clinics, etc., are also common. Ten prisoners became seriously ill in 2009 as a result of E. histolytica infection due to drinking contaminated water
in a city jail in northern Mindanao, Philippines.57 An epidemiological survey in Japan (1989) reported
that 20% of patients in a mental institution were infected with E. histolytica.58 Serious illness in a psychiatric clinic in Kaohsiung County in 2007 was reported to be due to E. histolytica infection.59 Outbreaks
of diarrheal disease due to E. histolytica in travelers returning from Thailand were reported in 1988.60
A total of 148 fatal cases of amoebiasis were reported by the Armed Forces Institute of Pathology during
1862–1953 in servicemen who returned from all over the world.61 At least 199 waterborne outbreaks of
human diseases due to parasitic protozoa were reported during the time period from 2004 to 2010, and
E. histolytica contributed to 9.4% of them.62
Improper handling of food or food products by Entamoeba-infected person is another reason for transmission of the diseases.63 In some developed countries, increased occurrence of amoebiasis has been
reported among men who have sex with men (MSM) and or those engage in oral–anal sex.15 It is also
reported that sexually active HIV-infected MSM are at greater risk of developing amoebiasis caused by
E. histolytica than the general population.64
Biology of Foodborne Parasites
8.5.2 Molecular Epidemiology
Genetic variation among E. histolytica isolates has been investigated extensively, as it may provide
important clues as to why most infections are asymptomatic, whereas some cause invasive disease.
Specific determinants for this diverse clinical manifestation are mostly unknown, but host genetics and
parasite genotypes both probably play significant roles.65,66 Proper identification and genetic characterization of clinical isolates have been proposed as a fundamental strategy to explore the relationship
between parasite genotypes and its virulence capacity. Genotyping Based on Conventional Polymorphic Markers
Some PCR-based genotyping studies utilized SREHP and chitinase (CHI) genes as polymorphic markers.67,68 A geographic diversity was detected among genotypes of field E. histolytica isolates, but no
association was seen between the clinical symptoms and the SREHP genotype at either the nucleotide or
predicted amino acid level.67 There was some evidence suggesting the existence of a bottleneck (demographic sweep) in population genetics and transcontinental spread of E. histolytica. These studies have
also demonstrated the usefulness of these alleles for distinguishing clinical isolates of E. histolytica and
E. dispar.68
The effectiveness of some other polymorphic loci for simultaneous differentiation and typing of
E. histolytica and E. dispar has been reported.69 Hsp1, Hsp2, Hsp5, and hsp6 and Dsp1, Dsp2, Dsp5, and
Dsp6 have been used as polymorphic markers for differentiation and typing of these two morphologically identical parasites. However, upon completion of the E. histolytica genome project, single locus
genotyping studies based on these conventional markers have become obsolete due to their low resolution. In contrast, multilocus genotyping systems based on highly polymorphic noncoding regions like
tRNA-linked STR loci have increased significantly typing resolution.70 tRNA-Linked STR Loci and Its Association with Virulence
Genome sequencing of E. histolytica identified as many as 4500 copies of tandemly arrayed tRNA
genes (10% of whole genome) within its genome. The intergenic regions of these tRNA genes consist of
STR sequences, which resemble the micro/minisatellites of eukaryotic genomes. The only difference is
that unlike randomly dispersed micro/minisatellites, STRs form a part of a larger unit, which itself is
tandemly arrayed. tRNA genes are thought to be “hot spots” for recombination, probably due to the formation of replication fork barriers by RNA polymerase III transcription complex. The intergenic tRNA
regions are A+T rich, resulting in frequent tandem duplication.35 Moreover, the average numbers of STRs
in the intergenic tRNA regions remained constant in E. histolytica for several years for a particular strain
in a continuous culture.70 These features make it a very useful tool for the quantification of evolutionary
divergence of this fascinating parasite as well as to explore the relationship between parasite genotype
and its virulence.
Several genotyping studies of E. histolytica based on these STR loci have been conducted by research
groups in different parts of the world. Ali et al.65 first reported the evidence for a possible link between
parasite genotype and outcome of infection with E. histolytica. E. histolytica clinical isolates obtained
from different disease outcome were genetically characterized using six tRNA-linked STR markers.
They identified a few genotypes that were significantly associated with diseases comparing to others. In a more recent study, a link between sequence type of tRNA-linked R–R locus and outcome of
E. ­histolytica infection has been identified.71 Sequence type 5RR was mostly associated with asymptomatic infections, whereas sequence type 10RR was predominantly associated with symptomatic (diarrhea/
dysentery and LA) infections. Avirulent E. histolytica strain with unique tRNA-linked short tandem
repeat patterns has also been found.47 These tRNA-linked STR loci have also been used to compare
genotypes of E. histolytica between stool- and liver abscess–derived samples from the same patients.
It has been reported that intestinal and liver abscess amoebae are genetically distinct.50
However, there are also some opposite observations. E. histolytica strains isolated from ALA patient
were genetically similar to those from asymptomatic patients.72 In a recent genotyping study based on
STR loci, E. histolytica isolates from asymptomatic patients are genetically closer to those from patients
with liver abscess than those from patients with diarrhea.73
Worldwide genealogy analysis of E. histolytica based on two tRNA-linked STR loci (DA-H and NK2)
has shown the existence of genetic recombination in this fascinating parasite.74 Nonrepetitive Coding Loci Significantly Linked with Parasite Virulence
In addition to these highly polymorphic repetitive loci, some nonrepetitive coding and noncoding regions
of E. histolytica have also been used in characterizations of clinical isolates with different infection outcomes.75 It was shown that polymorphism rates from coding and noncoding regions were significantly
different. This might be the result from variable selection pressures on these markers. Single nucleotide
polymorphism (SNP) within lectin (hgl3) gene was significantly associated with asymptomatic infections, indicating that lectin may be an important virulence factor of Entamoeba.
Whole genome sequencing of E. histolytica clinical isolates has shown that two SNPs within the
Cylicin-2 gene (EHI_080100/XM_001914351) are significantly associated with asymptomatic and liver
abscess outcomes.76 Therefore, tRNA-linked STR loci are probably surrogate markers for determination
of disease outcomes, whereas specific SNPs (nonsynonymous) identified within nonrepetitive regions
may be directly linked to parasite virulence.
Whole genome sequencing of different cultured lines of E. histolytica has shown big differences in
gene copy numbers.43 Moreover, it has provided evidence indicating that recombination has occurred
in the history of the sequenced genomes, suggesting that E. histolytica may reproduce sexually.43 This
information opens a new horizon in molecular epidemiology of amoebiasis. Transposable Elements and Genomic Rearrangements
Genome rearrangement or reorganization associated with invasive disease has also been reported in
E. histolytica.38 Transposons and repetitive DNA, which are abundantly present in Entamoeba, may assist
genome rearrangements. A comprehensive study of the repetitive elements of three Entamoeba genomes
identified hundred copies of class I transposons, that is, LINE and SINE elements as well as Entamoebaspecific repeats.77 These Entamoeba-specific ERE1 and ERE2 sequences represent a large proportion
of the genome of E. histolytica. The ERE2 sequence may be unique to E. histolytica, as it was found in
neither E. dispar nor E. invadens.77 Class II transposons (DNA transposons) are rare in E. histolytica or
E. dispar, but much more prevalent in E. invadens and E. moshkovskii.78 These results suggest expansion
and contraction of the number of transposable elements in different lineages, with likely consequences
for genome rearrangement. Comparisons of the genomes of E. histolytica and E. dispar have shown that
transposons have been active since these species diverged.77 Active transposition may still be occurring.79
A number of putative recent transpositions of EhSINE1 elements in the HM1/IMSS genome have been
identified.79 An extensive polymorphism of SINE occupancy among different strains of E. histolytica and
the genomic distribution of SINEs proved to be a valid method for typing of E. histolytica strains.80 Gene Families and Diversity
Possession of large gene families often indicates the importance and complexity of particular processes. E. histolytica contains a number of large multigene families.81 A large gene family encodes a
group of AIG1-like GTPases.81 Their precise function is unknown, but differential expression suggests
that they may be associated with virulence and/or adaptation to the intestinal environment.82 Members
of the AIG1-like family, among a number of gene families, often locate near transposons.81 Another
large gene family encodes proteins homologous to a bacterial fibronectin-binding protein (BspA of
Bacteroides forsythus), which encodes a large number (75–116) of proteins containing leucine-rich
repeats.81,82 Entamoeba encodes a very large number of Rab GTPases. These genes control vesicular
trafficking in the cell, and the size of the gene family points to the importance and complexity of
these processes in Entamoeba. In E. histolytica, 102 Rab GTPases, forming over 16 subfamilies, have
been annotated. Families encoding heavy- and light-chain subunits of the virulence factor Gal/GalNAc
Biology of Foodborne Parasites
lectin occur in multiple Entamoeba species, but the Gal/GalNAc lectin intermediate-chain subunit
genes have not been detected in species other than E. histolytica and E. dispar.10 The cysteine protease
family occurs in both E. histolytica and E. dispar,83 but the key virulence factor cysteine protease-5 is
a pseudogene in E. dispar.84 Southern blot evidence indicates that the Ariel surface proteins in E. histolytica are not present, or are highly divergent, in E. dispar.85
Epidemiology helps us to identify factors that influence disease distribution and determinants of disease occurrence in human populations in time and place, as well as factors related to disease transmission, manifestations, and outcome. Even though great advances have certainly been made regarding
the biological knowledge of E. histolytica, large gaps in the molecular epidemiology of amoebiasis in
different endemic areas remain. Multiple Infections and the Need for Sequencing
Mixed infections with both E. histolytica and E. dispar are possible. Infections of E. moshkovskii
together with E. dispar and/or E. histolytica have also been reported.86 PCR diagnosis using speciesspecific primers or PCR analysis of genetic markers such as chitinase, SREHP, and tRNA-linked STRs,
followed by sequencing, is essential to identify these mixed infections. These are now common practices.
8.6 Pathogenesis and Clinical Features
E. histolytica invades tissues and causes clinical disease through a sequence of events. The amoebic trophozoite first adheres to the colonic mucus and epithelial cells through interaction of Gal/GalNAc inhibitable lectin with host glycoconjugates. It next secretes proteolytic enzymes that efface the brush border
microvilli of enteric cells, disrupt the intestinal mucus and epithelial barrier, and facilitate tissue penetration. The trophozoite then kills the host epithelial and immune cells by amoebapore, a contact-dependent
cytolysin, causing characteristic flask-shaped ulcers. Finally, the parasite resists the host immune response
and survives to cause prolonged extraintestinal infection such as ALA. Trophozoites from stools of many
invasive patients contain ingested erythrocytes and have a much higher rate of erythrophagocytosis than
healthy human carriers. A phagocytosis-deficient mutant of E. histolytica has been isolated by Orozco
et al. (1983).87 This mutant apart from being poor in phagocytosis is also found to be low in virulence.
Thus, there seems to be a correlation between phagocytosis and virulence.
8.6.1 Clinical Features Symptomatic Infection
Depending on the affected organ, the clinical manifestations of amoebiasis are intestinal or extraintestinal. Intestinal Infections
Four clinical forms of invasive intestinal amoebiasis are known, all of which are generally acute. These
include dysentery or bloody diarrhea, fulminating colitis, amoebic appendicitis, and amoeboma of the
Dysenteric and diarrheic syndromes account for 90% of cases of invasive intestinal amoebiasis.
Patients with dysentery have an average of three to five mucosanguineous evacuations per day, with
moderate colic pain preceding discharge, and have rectal tenesmus. In patients with bloody diarrhea,
although evacuations are few, the stools contain blood. Fever and systemic manifestations are generally
absent. These syndromes constitute the classic ambulatory dysentery and can easily be distinguished
from that of bacterial origin, where the patient frequently complains of systemic signs and symptoms
such as fever, chills, headache, malaise, anorexia, vomiting, cramping abdominal pain, and tenesmus.20
Amoebic colitis (70% of cases) is usually associated with segmentary ulceration of the colon. If ulceration with perforation occurs, it occurs mainly in the cecum.88 As the ulcer deepens and progresses,
it forms the classic flask-shaped ulcer of amoebic colitis, which extends from the mucosa and muscularis
mucosa into the submucosa. The patient presents with complaints of severe abdominal pain, intense and
unrelenting tenesmus, and 20 or more episodes of bloody diarrhea within a 24 h period. Other symptoms
include nausea, anorexia, fever, tachycardia, and hypotension.
Amoebic appendicitis constitutes about 1% of all cases of acute appendicitis in adults living in areas
where E. histolytica is endemic. In amoebic appendicitis, the parasite invades the ileocecal appendix
resulting in inflammation, necrosis, and eventually perforation. Symptoms include pain, guarding in the
lower right abdominal quadrant, fever, tachycardia, nausea, and vomiting. When the cecum is involved,
a mucosanguineous diarrhea may also be present. Amoeboma, the result of chronic ulceration, is predominantly found in the cecum, sigmoid colon, and rectum. The patient presents with painful, palpable
abdominal mass and bloody dysentery. Extraintestinal Infection
ALA is the most common extraintestinal manifestation of E. histolytica infection. Liver abscess can occur
concurrently with colitis, and it is estimated that 10% of individuals with amoebic colitis will develop
an ALA.88 The most common clinical symptoms include fever, chills, nausea, weakness, malaise, and a
constant, dull, aching abdominal pain in the right upper quadrant or epigastrum.89 The patient experiences
right pleuritic pain or shoulder pain if the diaphragmatic surface of the liver is involved. Other symptoms
include weight loss and myalgia. In rare cases, a large ALA compresses the biliary ductal system, resulting
in obstructive jaundice. Hepatomegaly with pain on palpation is one of the most important clinical signs
for ALA. Point tenderness over the liver, or below the ribs, or in the intercostal spaces is a typical finding.88
The respiratory tract, brain, and urinary, genital, and rectal tracts may also be affected. Patients
develop coughing, pleuritic chest pain, respiratory distress and could have pleuropulmonary amoebiasis, secondary to amoebic liver abscess rupture through the diaphragm. This complication occurs in
7%–20% of patients with ALA and with resultant empyema and effects on the lung parenchyma, occasionally leading to an erroneous diagnosis of bacterial pneumonia. Amoebic brain abscesses are very
rare and arise almost exclusively alongside amoebic liver abscess (<0.1% of liver abscess cases). Sudden
onset of symptoms (headache, vomiting, seizures, and mental status changes) and rapid progression to
death have been reported.90 Asymptomatic Infection
Noninvasive intestinal infection is the most common presentation of E. dispar infection. Patients may
have some ill-defined gastrointestinal complaints, but for the most part tolerate the infection well. The
vast majority of E. histolytica infections are asymptomatic. Either these are caused by nonpathogenic
strains of the parasite, in contrast to invasive infection resulting from pathogenic strains, or host factors
decide the course of infection even if all E. histolytica strains are pathogenic. From the limited information available so far, it appears that the property of pathogenesis is determined more by quantitative
levels of key molecules than by the total absence of these in nonpathogenic species.
8.7 Treatment and Prevention
8.7.1 Treatment
Dysentery, if associated with infection, is initially managed by maintaining fluid intake using oral rehydration. In ideal situations, no antimicrobial therapy should be administered until microscopy or other
confirmatory diagnosis establishes the specific infection involved. When laboratory services are not
available, it may be necessary to administer a combination of drugs, including an amoebicidal drug to
kill the parasite and an antibiotic to treat any associated bacterial infection.86
Amoebic dysentery usually calls for a two-pronged attack. Treatment should start with a 10-day
course of the antimicrobial drug metronidazole (Flagyl). To kill the parasite, the physician can
Biology of Foodborne Parasites
prescribe a course of diloxanide furoate (available only through the Centers for Disease Control and
Prevention), paromomycin (Humatin), or iodoquinol (Yodoxin).
The main objective of amoebicidal drugs is to eliminate E. histolytica from the intestinal lumen, intestinal wall, and the hepatic tissues. Invasive amoebiasis should be treated with drugs like emetine, dehydroemetine, chloroquine, metronidazole, and tinidazole, as they act within the intestinal lumen as well
as in the liver.86 For the treatment of mild to moderate infection, a mixture of tetracycline with “luminal”
amoebicide and chloroquine is useful. For severe infections, treatment should include a mixture of dehydroemetine combined with tetracycline and a “luminal” amoebicide. A treatment with emetine hydrochloride and emetine bismuth iodide proves to be beneficial for the treatment of amoebic dysentery. For
the treatment of ALA, metronidazole should be the drug of choice since it accumulates the most in liver
and is highly effective for eliminating trophozoite present in the hepatic tissues.86,91
Emetine is one of the best drugs for ALA treatment. The drug found in Ipecac or ipecacuanha as an
alkaloid–emetine and cephaline acts directly on the trophozoites. Its concentration and activity are both
higher in the liver than in the intestine; therefore, emetine is more effective for treating liver abscess
caused by E. histolytica. The side effect of this drug is prominent and includes muscular tremors, weakness, and pain in the extremities as it is detoxified and eliminated slowly. Nausea, vomiting, and bloody
diarrhea are also reported in certain cases. Cardiac toxicity is reported in emetine treatment, but this
is very rare in normal individuals. Anemic patients are said to have a higher incidence of heart damage than normal persons. It is advisable that patients undergoing emetine treatment should have regular
checks on blood pressure and ECG and bed rest during and after the treatment. Emetine is administered
intramuscularly or subcutaneously at a dose less than 650 mg or 10 mg/kg body weight. After 2–6 weeks,
treatment could be repeated when liver abscess relapses.
Antibiotics such as tetracycline, chlortetracycline, oxytetracycline, and erythromycin are sometimes
used as indirect amoebicides. They are effective in the intestinal lumen but have no activities in the
hepatic tissues, thus are not used for treatment of hepatic abscesses.
8.7.2 Prevention
Good sanitary practice and proper sewage disposal or treatment are necessary for the prevention of
E. histolytica infection at the community level. Hands must be washed thoroughly with hot running
water and soap for at least 10 s before handling and consumption of food, after changing diapers, or
using the toilet. Eating raw vegetables, fruits, and salads in endemic areas should be avoided, as they
may have been fertilized by human feces. Drinking water should be boiled or treated with iodine tablets.
E. histolytica cysts are usually resistant to chlorination; therefore, filtration of water supplies is necessary
to reduce the incidence of infection.89
As E. histolytica only infects humans and some higher nonhuman primates, a large animal reservoir
is therefore absent, making it theoretically possible to eliminate amoebiasis from the world by a vaccine.
Recently, a Gal-lectin-based intranasal synthetic peptide vaccine was evaluated in a primate model of
E. histolytica intestinal infection; it was highly efficacious in preventing experimental E. histolytica
infection and colitis in baboons.92
1. World Health Organization. 1998. Executive summary, The World Health Report 1998. Life in the
21st century—A vision for all. World Health Organization, Geneva, Switzerland.
whr/1998/en/whr98_en.pdf (accessed November 25, 2014).
2. Cox, D. R. Regression models and life tables. J. R. Stat. Soc. B 34, 187–220, 1972.
3. Kiple, K. Cambridge World History of Human Diseases. Cambridge University Press, Cambridge, U.K., 1993.
4. Crosby, A. W. Ecological Imperialism: The Biological Expansion of Europe, 900–1900. Cambridge
University Press, Cambridge, U.K., 1986.
5. Clark, C. G. et al. Molecular systematics of the intestinal amoebae. In Evolutionary Relationship
among Protozoa, Coombs, G. H., Vickerman, K., Sleigh. M. A., and Warren. A., eds. Kluwer Academic
Publishers, Dordrecht, the Netherland, pp. 169–180, 1998.
6. Dobell, C. Researches on the intestinal protozoa of monkeys and man. Parasitology 20, 357–412, 1928.
7. Ghosh, S. K. et al. Chitinase secretion by encysting Entamoeba invadens and transfected E. histolytica
trophozoites: Localization of secretory vesicles, ER and Golgi apparatus. Infect. Immun. 67, 3073–3081,
8. Mai, Z. et al. Hsp60 is targeted to a cryptic mitochondrion derived organelle (“crypton”) in the microaerophilic protozoan parasite Entamoeba histolytica. Mol. Cell Biol. 19, 2198–2205, 1999.
9. Tovar, J., Fischer, A., and Clark, C. G. The mitosome, a novel organelle related to mitochondria in the
amitochondrial parasite Entamoeba histolytica. Mol. Microbiol. 32, 1013–1021, 1999.
10. Clark, C. G. et al. Structure and content of the Entamoeba histolytica genome. Adv. Parasitol. 65, 51–190,
11. Edman, U., Meza, I., and Agabian, N. Genomic and cDNA actin sequences from a virulent strain of
Entamoeba histolytica. Proc. Natl. Acad. Sci. USA 84, 3024–3028, 1987.
12. Katiyar, S. K. and Edlind, T. Entamoeba histolytica encodes a highly divergent b tubulin. J. Euk.
Microbiol. 43, 31–34, 1996.
13. Vargas, M. et al. Molecular characterization of myosin IB from the lower eukaryote Entamoeba histolytica, a human parasite. Mol. Biochem. Parasitol. 86, 61–73, 1997.
14. Flores-Robles, D. et al. Entamoeba histolytica: A b1 integrin-like fibronectin receptor assembles a signaling complex similar to those of mammalian cells. Exp. Parasitol. 103, 8–15, 2003.
15. Hung, C. C., Chang, S. Y., and Ji, D. D. Entamoeba histolytica infection in men who have sex with men.
Lancet Infect. Dis. 12(9), 729–736, 2012.
16. Arroyo-Begovich, A., Cárabez-Trejo, A., and Ruíz-Herrera, J. Identification of the structural component
in the cyst wall of Entamoeba invadens. J. Parasitol. 66, 735–741, 1980.
17. Debnath, A. Studies on differentially expressed genes in Entamoeba histolytica during human collagen
type I and Ca2+ activation. PhD dissertation, Calcutta University, Kolkata, India, 2003.
18. Heredia, R. D., Fonseca, J. A., and López, M. C. Entamoeba moshkovskii perspectives of a new agent to
be considered in the diagnosis of amebiasis. Acta Trop. 123, 139–145, 2012.
19. Makioka, A. et al. Expression analysis of Entamoeba invadens profilins in encystations and excystation.
Parasitol. Res. 110(6), 2095–2104, 2012.
20. Stanley, S. L. Amoebiasis. Lancet 361, 1025–1034, 2003.
21. Picazarri, K., Nakada-Tsukui, K., and Nozaki, T. Autophagy during proliferation and encystation in the
protozoan parasite Entamoeba invadens. Infect. Immun. 76, 278–288, 2008.
22. Tovy, A., Tov, R. S., Gaentzsch, R., Helm, M., and Ankri, S. A new nuclear function of the Entamoeba
histolytica glycolytic enzyme enolase: The metabolic regulation of cytosine-5 methyltransferase 2
(Dnmt2) activity. PLoS Path., 6(2), 1–13, 2010.
23. Serrano, R. and Reeves, R. E. Glucose transport in Entamoeba histolytica. Biochem. J. 144(1), 43–48,
24. Takeuchi, T., Weinbach, E. C., and Diamond, L. S. Entamoeba histolytica: Localization and characterization of phosphoglucomutase, uridine diphosphate glucose pyrophosphorylase, and glycogen synthase.
Exp. Parasitol. 43(1), 115–121, 1977.
25. Bragg, P. D. and Reeves, R. E. Pathways of glucose dissimilation in the Laredo strain of Entamoeba
histolytica. Exp. Parasitol. 12(5), 393–400, 1962.
26. Ghulam, J. et al. Metabolic profiling of the protozoan parasite Entamoeba invadens revealed activation
of unpredicted pathway during encystation. PLoS ONE 10, 1371, 2007.
27. Yi, D., Lee, R. T., Longo, P., Boger, E. T., Lee, Y. C., Petri, W. A. Jr, and Schnaar, R. L. Substructural
specificity and polyvalent carbohydrate recognition by the Entamoeba histolytica and rat hepatic N-acetyl
galactosamine/galactose lectins. Glycobiology 8(10), 1037–1043, 1998.
28. Williams, K., Lowe, P. N., and Leadlay, P. F. Purification and characterization of pyruvate: Ferredoxin
oxidoreductase from the anaerobic protozoon Trichomonas vaginalis. Biochem. J. 246(2), 529–536,
29. Bruchhaus, I. and Tannich, E. Induction of the iron-containing superoxide dismutase in Entamoeba histolytica by a superoxide anion-generating system or by iron chelation. Mol. Biochem. Parasitol. 67(2),
281–288, 1994.
30. Jeelani, G., Sato, D., Husain, A., Escueta-de Cadiz, A., Sugimoto, M., Soga, T., Suematsu, M., and
Nozaki, T. Metabolic profiling of the protozoan parasite Entamoeba invadens revealed activation of
unpredicted pathway during encystation. PLoS ONE 7(5), e37740, 2012.
Biology of Foodborne Parasites
31. Gelderman, A. H., Keister, D. B., Bartgis, I. L., and Diamond, L.S. Characterization of the deoxyribonucleic acid of representative strains of Entamoeba histolytica, E. histoytica-like amebae, and E. moshkovskii. J. Parasitol. 57(4), 906–911, 1971.
32. Mukherjee, C., Clark, C.G., and Lohia, A. Entamoeba shows reversible variation in ploidy under different
growth conditions and between life cycle phases. PLoS Negl. Trop. Dis. 2, e281, 2008.
33. Willhoeft, U. and Tannich, E. Fluorescence microscopy and fluorescence in situ hybridization of
Entamoeba histolytica nuclei to analyse mitosis and the localization of repetitive DNA. Mol. Biochem.
Parasitol. 105, 291–296, 2000.
34. Loftus, B. et al. The genome of the protist parasite Entamoeba histolytica. Nature 433, 865–868, 2005.
35. Tawari, B., Ali, I. K. M., Scott, C., Quail, M. A., Berriman, M., Hall, N., and Clark, C. G. Patterns
of evolution in the unique tRNA gene arrays of the genus Entamoeba. Mol. Biol. Evol. 25, 187–198,
36. Cheng, X. J. et al. Intermediate subunit of the Gal/GalNAc lectin of Entamoeba histolytica is a member
of a gene family containing multiple CXXC sequence motifs. Infect Immun. 69(9), 5892–5898, 2001.
37. Clark, C. G., Ali, I. K. M., Zaki, M. B., Loftus, J., and Hall, N. Unique organization of tRNA genes in
Entamoeba histolytica. Mol. Biochem. Parasitol. 146, 24–29, 2006.
38. Ali et al. Tissue invasion by Entamoeba histolytica: Evidence of genetic selection and/or DNA reorganization events in organ tropism. PLoS Negl. Trop. Dis. 2(4), 219, 2008.
39. Dellen et al. LINEs and SINE-like elements of the protist Entamoeba histolytica. Gene 297(2002), 229–239,
40. Tanyuksel, M. and Petri, W. A. Jr. Laboratory diagnosis of amebiasis. Clin. Microbiol. Rev. 16(4), 713–
729, 2003.
41. Chawla, T. C., Irshad, M., Gandhi, B. M., and Tandon, B. N. Immunological characterization of axenic
Entamoeba histolytica related antigens. Indian J. Med. Res. 86, 457–461, 1987.
42. Abe, N. et al. Entamoeba histolytica outbreaks in institutions for the mentally retarded. Jpn. J. Infect.
Dis. 52, 135–136, 1999.
43. Weedall, G. D. and Hall, N. Evolutionary genomics of Entamoeba. Res. Microbiol. 162, 637–645, 2011.
44. Ayeh-Kumi, P. F., Ali, I. M., Lockhart, L. A., Gilchrist, C. A., Petri, W. A. Jr, and Haque, R. Entamoeba
histolytica: Genetic diversity of clinical isolates from Bangladesh as demonstrated by polymorphisms in
the serine-rich gene. Exp. Parasitol. 99(2), 80–88, 2001.
45. Clark, C. G. and Diamond, L. S. Entamoeba histolytica: A method for isolate identification. Exp.
Parasitol. 77, 450–455, 1993.
46. Haghighi, A. S. et al. Remarkable genetic polymorphism among Entamoeba histolytica isolates from a
limited geographic area. J. Clin. Microbiol. 40, 4081–4090, 2002.
47. Escuta-de, C. A. et al. Identification of an avirulent Entamoeba histolytica strain with unique tRNAlinked short tandem repeat markers. Parasitol. Int. 59, 75–81, 2010.
48. Vega, H. et al. Cloning and expression of chitinases of Entamoeba. Mol. Biochem. Parasitol. 85, 139–
147, 1997.
49. Beck, D. L. et al. Entamoeba histolytica: Sequence conservation of the Gal/GalNAc lectin from clinical
isolates. Exp. Parasitol. 101, 157–163, 2002.
50. Ali, I. M., Clark, C. G., and Petri, W. A. Jr. Molecular epidemiology of amebiasis. Infect. Genet. Evol.
8(5), 698–707, 2008.
51. Haque, R. et al. Amoebiasis. N. Eng. J. Med. 348, 1565–1573, 2003.
52. Parija, S. C. and Khairnar, K. Entamoeba moshkovskii and Entamoeba dispar associated infections in
Pondicherry, India. J. Health Pop. Nutr. 23, 292–295, 2005.
53. Mukherjee, A. K. et al. Hospital based surveillance of enteric parasites in Kolkata. BMC Res. Notes 2,
110, 2009.
54. Clark, C. G. and Diamond, L. S. Intraspecific variation and phylogenetic relationships in the genus
Entamoeba as revealed by riboprinting. J. Eukaryot. Microbiol. 44, 142–154, 1997.
55. Markell, E. K. The 1933 Chicago outbreak of amebiasis. West. J. Med. 144(6), 750, 1986.
56. Barwick, R. S. et al. Outbreak of amebiasis in Tbilisi, Republic of Georgia 1998. Am. J. Trop. Med. Hyg.
67(6), 623–631, 2002.
57. Herriman, R. 2009. Amoebiasis outbreak in Philippine jail. (accessed November 25, 2014).
58. Nagakura, K. et al. An outbreak of amebiasis in an institution for the mentally retarded in Japan. Jpn. J.
Med. Sci. Biol. 42(2), 63–76, 1989.
59. Chen, Y.-H., Hung, M.-N., Chan, Y.-H., Yeh, H.-C., Chen, M.-J., Lin, L.-J., Chiou, H.-Y., Chang, C.-C.,
Lee, Y.-S., and Lin, C.-W. Outbreak of Amoebiasis in a psychiatric hospital-KaoHsiung County. Taiwan
Epi. Bull. 24(10), 753–765, 2008.
60. de Lalla, F., Rizzardini, G., Cairoli, G. A., Rinaldi, E., Santoro, D., and Ostinelli, A. Outbreak of amoebiasis in tourists returning from Thailand. Lancet 2(8615), 847, 1988.
61. Kean, B. H., Gilmore, H. R. Jr, and Van Stone, W. W. Fatal amebiasis: Report of 148 fatal cases from the
armed forces institutes of pathology. Ann. Intern. Med. 44(5), 831–843, 1956.
62. Baldursson, S. and Karanis, P. Waterborne transmission of protozoan parasites: Review of worldwide
outbreaks—An update 2004–2010. Water Res. 45, 6603–6614, 2011.
63. Ben Ayed, S., Ben, A. R., Mousli, M., Aoun, K., Thellier, M., and Bouratbine, A. Molecular differentiation of Entamoeba histolytica and Entamoeba dispar from Tunisian food handlers with amoeba infection
initially diagnosed by microscopy. Parasite 15(1), 65–68, 2008.
64. James, R., Barratt, J., Marriott, D., Harkness, J., and Stark, D. Seroprevalence of Entamoeba histolytica
infection among men who have sex with men in Sydney, Australia. Am. J. Trop. Med. Hyg. 83(4), 914–
916, 2010.
65. Ali, I. K. M., Mondal, U., Roy, S., Haque, R., Petri, W. A. Jr., and Clark, C. G. Evidence for a link between
parasite genotype and outcome of infection with Entamoeba histolytica. J. Clin. Microbiol. 45, 285–289, 2007.
66. Duggal, P. R., Haque, R., Roy, S., Mondal, D., Sack, R. B., and Farr, B. M. Influence of human Leukocyte
antigen class II alleles on susceptibility to Entamoeba histolytica infection in Bangladeshi children.
J. Infect. Dis. 189, 520–526, 2004.
67. Haghighi, A., Kobayashi, S., Takeuchi, T., Thammapalerd, N., and Nozaki, T. Geographic diversity
among genotypes of Entamoeba histolytica field isolates. J. Clin. Microbiol. 41, 3748–3756, 2003.
68. Ghosh, S., Frisardi, M., Ramirez-Avila, L., Descoteaux, S., Sturm-Ramirez, K., Newton-Sanchez, O. A.,
Santos-Preciado, J. I., Ganguly, C., Lohia, A., Reed, S., and Samuelson, J. Molecular epidemiology of
Entamoeba spp.: Evidence of a bottleneck (demographic sweep) and transcontinental spread of diploid
parasites. J. Clin. Microbiol. 38, 3815–3821, 2000.
69. Zaki, M., Meelu, P., Sun, W., and Clark, C. G. Simultaneous differentiation and typing of Entamoeba
histolytica and Entamoeba dispar. J. Clin. Microbiol. 40, 1271–1276, 2002.
70. Ali, I. K. M., Zaki, M., and Clark, C. G. Use of PCR amplification of tRNA gene-linked short tandem
repeats for genotyping Entamoeba histolytica. J. Clin. Microbiol. 43, 5842–5847, 2005.
71. Ali, I. K. M., Haque, R., Alam, F., Kabir, M., Siddique, A., and Petri, W. A. Evidence for a link between
locus R-R sequence type and outcome of infection with Entamoeba histolytica. Clin. Microbiol. Infect.
18, 235–237, 2012.
72. Feng, M., Cai, J., Yang, B., Fu, Y., Min, X., Tachibana, H., and Cheng, X. Unique short tandem repeat
nucleotide sequences in Entamoeba histolytica isolates from China. Parasitol. Res. 111, 1137–1142, 2012.
73. Das, K., Mukherjee, A. K., Chowdhury, P., Sehgal, R., Bhattacharya, M. K., Hashimoto, T., Nozaki,
T., and Ganguly, S. Multilocus sequence typing system (MLST) reveals a significant association of
Entamoeba histolytica genetic patterns with disease outcome. Parasitol. Int. 63, 308–314, 2014.
74. Zermeno, V., Ximenez, C., Moran, P., Valadez, A., Valenzuela, O., and Rascon, E. Worldwide genealogy of Entamoeba histolytica: An overview to understand haplotype distribution and infection outcome.
Infect. Genet. Evol. 17, 243–252, 2013.
75. Bhattacharya, D., Haque, R., and Singh, U. Coding and noncoding genomic regions of Entamoeba histolytica have significantly different rates of sequence polymorphisms: Implications for epidemiological
studies. J. Clin. Microbiol. 43, 4815–4819, 2005.
76. Gilchrist, C. A. et al. A multilocus sequence typing system (MLST) reveals a high level of diversity and
a genetic component to Entamoeba histolytica virulence. BMC Microbiol. 12, 151, 2012.
77. Lorenzi, H., Thiagarajan, M., Haas, B., Wortman, J., Hall, N., and Caler, E. Genome wide survey, discovery and evolution of repetitive elements in three Entamoeba species. BMC Genomics 9, 595, 2008.
78. Pritham, E. J., Feschotte, C., and Wessler, S. R. Unexpected diversity and differential success of DNA
transposons in four species of Entamoeba protozoans. Mol. Biol. Evol. 22, 1751–1763, 2005.
79. Huntley, D. M., Pandis, I., Butcher, S. A., and Ackers, J. P. Bioinformatic analysis of Entamoeba histolytica SINE1 elements. BMC Genomics 11, 321, 2010.
Biology of Foodborne Parasites
80. Kumari, V. et al. Genomic distribution of SINEs in Entamoeba histolytica strains: Implication for genotyping. BMC Genomics 14, 432, 2013.
81. Lorenzi, H. A., Puiu, D., Miller, J. R., Brinkac, L. M., Amedeo, P., Hall, N., and Caler, E. V. New assembly, reannotation and analysis of the Entamoeba histolytica genome reveal new genomic features and
protein content information. PLoS Negl. Trop. Dis. 4, e716, 2010.
82. Davis, P. H., Schulze, J., and Stanley, S.L. Jr. Transcriptomic comparison of two Entamoeba histolytica
strains with defined virulence phenotypes identifies new virulence factor candidates and key differences
in the expression patterns of cysteine proteases, lectin light chains, and calmodulin. Mol. Biochem.
Parasitol. 151, 118–128, 2007.
83. Bruchhaus, I., Jacobs, T., Leippe, M., and Tannich, E. Entamoeba histolytica and Entamoeba dispar:
Differences in numbers and expression of cysteine proteinase genes. Mol. Microbiol. 22, 255–263, 1996.
84. Willhoeft, U., Hamann, L., and Tannich, E. A DNA sequence corresponding to the gene encoding cysteine proteinase 5 in Entamoeba histolytica is present and positionally conserved but highly degenerated in
Entamoeba dispar. Infect. Immun. 67, 5925–5929, 1999b.
85. Willhoeft, U., Buss, H., and Tannich, E. DNA sequences corresponding to the ariel gene family of
Entamoeba histolytica are not present in E. dispar. Parasitol. Res. 85, 787–789, 1999a.
86. Gonzales, M. L., Dans, L. F., and Martinez, E. G. Antiamoebic drugs for treating amoebic colitis.
Cochrane Database Syst Rev. 15(2), 1–32, 2009.
87. Orozco, E., Guarneros, G., Martinez-Palomo, A., and Sánchez, T. Entamoeba histolytica. Phagocytosis
as a virulence factor. J. Exp. Med. 158(5), 1511–1521, 1983.
88. Stanley, S. L. Jr. and Reed, S. L. Entamoeba histolytica: Parasite-host interactions. Am. J. Physiol.
Gastrointest. Liver. Physiol. 280, G1049–G1054, 2001.
89. Petri, W. A. Jr. et al. Estimating the impact of amebiasis on health. Parasitol. Today 16, 320–321, 2000.
90. Ravdin, J. I. Amebiasis: Review. Clin. Infect. Dis. 20, 1453–1466, 1995.
91. Madigan, M. T. et al. Brock Biology of Microorganisms. Pearson Education, Inc., Upper Saddle River,
NJ, 2003.
92. Mohamed, D. A. et al. Efficacy of a Gal-lectin subunit vaccine against experimental Entamoeba histolytica infection and colitis in baboons (Papio sp.). Vaccine 30(20), 3068–3075, 2012.
Enterocytozoon bieneusi
Mónica Santín-Durán
Morphology and Classification..................................................................................................... 150
Biology, Genetics, and Genomics................................................................................................. 150
Diagnosis and Typing.................................................................................................................... 150
Epidemiology and Molecular Epidemiology.................................................................................151
9.5.1 E. bieneusi in Humans......................................................................................................163
9.5.2 E. bieneusi in Animals..................................................................................................... 164 Nonhuman Primates......................................................................................... 164 Companion Animals......................................................................................... 164 Livestock............................................................................................................165 Birds.................................................................................................................. 166 Other Animals.................................................................................................. 166
9.5.3 E. bieneusi in Water......................................................................................................... 166
9.5.4 E. bieneusi in Food............................................................................................................167
9.6 Pathogenesis and Clinical Features...............................................................................................167
9.7 Treatment and Prevention..............................................................................................................168
9.1 Introduction
Microsporidia is a diverse group of obligate intracellular parasites that includes more than 1200 named
species.1 Microsporidia has a wide range of vertebrates and invertebrate as hosts; most of the species infect
invertebrates and fish, but at least 14 species in 8 genera infect humans.2 The most common microsporidian
species infecting humans are Enterocytozoon bieneusi, Encephalitozoon cuniculi, Encephalitozoon intestinalis, and Encephalitozoon hellem. Of these four species, E. bieneusi is the most frequently diagnosed
species in humans worldwide, mainly associated with chronic diarrhea and wasting syndrome.3,4 It has
been recognized as an opportunistic pathogen of immunocompromised patients (HIV-infected people
or organ transplant recipients),5–10 but immunocompetent individuals have also been found infected.11–16
In addition, E. bieneusi has been identified in a variety of wild, farm, and companion mammals, raising
the question on the importance of animal reservoirs in the epidemiology of this parasite.17 E. bieneusi
has been detected in retail fresh food produce (raspberries, sprouts, and lettuce)18 in Poland and has been
found to be the cause of a large foodborne outbreak in Sweden.19 The identification of spores of E. bieneusi
in water supplies suggests that water can be a potential vehicle in the transmission of this parasite not only
for humans but also for animals.20 Identification of microsporidia in water sources has led to their inclusion on the National Institute of Allergy and Infectious Diseases Category B Priority List of biodefense
pathogens ( and in the
Environmental Protection Agency drinking water Contaminant Candidate List of concern for waterborne
transmission (
Biology of Foodborne Parasites
9.2 Morphology and Classification
The spore, the infectious stage, is the most characteristic form of the microsporidia and the only state
that is viable outside of a host cell. E. bieneusi spores are small (0.5 × 1.5 µm) and oval and surrounded
by an outer electrodense glycoprotein layer and electron-lucent endospore layer composed primarily of
chitin and an inner plasma membrane.21 Within the spore membrane is the sporoplasm (or cytoplasm)
of the spore, which is the infectious material.22 The sporoplasm contains in E. bieneusi a single nucleus
(monokaryon). Spores have a unique extrusion apparatus composed of an anchoring disk, a polar filament (or polar tube), and a polaroplast that allow them to penetrate and infect the host cell.23 The polar
filament is coiled within the spore approximately six times in E. bieneusi. The polar filament terminates
at the posterior vacuole. There is a lack of typical mitochondria, Golgi, and peroxisomes, and it possesses
small ribosomes like those of prokaryotes.
Microsporidia have been taxonomically problematic since its discovery. Taxonomy was based originally on morphology, ultrastructure, biology, and habitat features, but now molecular phylogenetics are
also applied for classification of this organisms. The initial analysis of the small subunit ribosomal
RNA genes indicated that Microsporidia may represent one of the earliest-diverging eukaryotic lineages.24 However, subsequent phylogenetic analysis using multiple genes supports a fungal origin for the
Microsporidia.25–27 E. bieneusi was first identified in 1985,28 when it was described and named as a new
genus and species.14 The new genus was established on the basis of its development in direct contact with
host cytoplasm, the precocious development of the spore organelles in the sporont, and the poor development of the endospore layer of the spore wall.29
9.3 Biology, Genetics, and Genomics
E. bieneusi is an obligate intracellular parasite and has no active stages outside host cells. The infectious
stage is the spore; spores are shed with feces and are relatively resistant to the environment. After a spore
is ingested by the right host, the coiled polar tube discharges, injecting the sporoplasm and nucleus into
the host cell. Once inside the host cell, the parasite is referred to as meront and begins a state of growth
and division. Young proliferative stages become elongate and undergo nuclear division. Proliferative
plasmodial cells contain multiple elongated nuclei. Following division, the extrusion apparatus (including the polar filament, polaroplast, and posterior vacuole) begins to develop. Sporogonial plasmodium
develops disks, some in stacks or arcs in stages of polar tube formation. Individual nuclei with polar tube
complexes segregate and mature into separate sporoblast cells that develop into mature spores. Spores
released from host cells are excreted with feces into the environment where they can be ingested by the
next host.
E. bieneusi has a reduced and compacted genome. The first large-scale genomic database showed
many traits associated with genome compaction including high gene density, short intergenic regions,
shortened protein-coding sequences, and few introns.30
9.4 Diagnosis and Typing
Microsporidia are difficult to diagnose even though significant progress has been made in the last decade.
Microscopic examination of microsporidian spores in feces is quite difficult because the spores are very
small (E. bieneusi spores are 0.5 × 1.5 µm); accurate diagnosis is highly dependent on the expertise
of the examiner. Furthermore, bright-field and fluorescence microscopic methods are not sufficient to
determine the species of Microsporidia. Transmission electron microscopy (TEM) has been the gold
standard to identify the polar filament and other species-specific ultrastructural characters necessary for
the identification of the microsporidian species. However, identification based on ultrastructural studies
using TEM is not feasible for routine diagnosis because it is expensive, time consuming, and labor intensive and requires expensive equipment and trained personal.
Enterocytozoon bieneusi
Light microscopy is an inexpensive method of diagnosing microsporidian infections even though
it does not allow identification of microsporidia to the species level. Microscopic diagnostic tests for
microsporidia have been reviewed in detail by Weber et al.31 The most widely used staining technique
is the modified trichrome stains or its adaptations (Weber Green or Ryan Blue). Stained spores are pink
against a blue/green background. This technique requires time to perform (about 90 min). A rapid hot
Gram-chromotrope staining method that cuts down the staining time to 5 min is available, where stained
spores are dark violet against a green/black background. Acid-fast trichrome stain has the advantage to
stain not only microsporidian spores but also Cryptosporidium oocysts at the same time. Using Giemsa
stain, spores (light blue) are difficult to differentiate from debris in stool smears, although in intestinal
biopsies their identification of spores is easier because of the presence of less debris. Chemofluorescent
optical brightening agents, such as Uvitex 2B, Fungi-Fluor, or Calcofluor white, stain chitin in the endospore layer of the spore. They are useful in the quick screening of spores in fecal smears. Stained spores
appear as white or turquoise oval halos under fluorescence microscopy. The occurrence of false positives
is the main problem because small fungi and some other small particles present in feces may also fluoresce. Immunofluorescent staining techniques using poly- and/or monoclonal antibodies have limited
availability and are only available in research laboratories.
The use of serologic assays is limited for E. bieneusi most likely because of the lack of in vitro cultivation methods and animal models. There are no serologic assays commercially available for E. bieneusi.
In the literature, two studies have determined the seropositivity for E. bieneusi in the Czech Republic
and Russia using indirect immunofluorescence antibody assay to detect specific antibodies against
E. bieneusi using purified whole E. bieneusi spores.32,33
PCR-based methods are commonly used in research laboratories for detecting E. bieneusi. They offer
the advantage of further analysis of the PCR products to identify species and genotypes. However, these
techniques are mainly used in research and some government public health laboratories, and their use
in clinical diagnostic laboratories is still limited. Molecular diagnostic tests for microsporidia have been
reviewed in detail by Ghosh and Weiss.34 Nucleic acids may be extracted from tissues or feces; an efficient DNA extraction method is needed, as the technique used to extract DNA for amplification can
affect the sensitivity of a PCR. Commercial extraction kits have shown their usefulness to get DNA
of good quality for PCR amplification. PCR amplification using stool specimens for the detection
of E. bieneusi could be difficult because of PCR inhibitors and the difficulty of spore disruption. PCR
methods applied for diagnostic commonly use primers that target rDNA genes. In addition to PCR-based
methods, fluorescent in situ hybridization–based methods have also been used to detect microsporidia.35
However, they are expensive, time consuming, and less sensitive than PCR.
Lower sensitivity of light microscopy in comparison with PCR has been shown in several studies.36–39
A blinded, externally controlled multicenter evaluation of light microscopy and PCR for detection of
microsporidia in stool specimens showed that PCR detected as few as 100 spores/g of stool, while detection limit of 10,000 spores/g of stool was shown by light microscopy.36 A spore shedding pattern study
of E. bieneusi in asymptomatic cases followed children in an orphanage in Bangkok, Thailand.38 Stool
examination was performed daily until they are found negative for E. bieneusi by Gram-chromotrope
staining for 2 months. PCR could detect E. bieneusi in the Gram-chromotrope-negative stool specimens
during and after the period of E. bieneusi positivity by light microscopy suggesting that light microscopy
may not be able to detect low levels of spore shedding.38 Similarly, a higher prevalence by PCR than by
Gram-chromotrope-staining-based microscopy was observed in pigs and humans in a survey of a farm
community in Central Thailand.39
9.5 Epidemiology and Molecular Epidemiology
E. bieneusi has a worldwide distribution and the ability to infect a wide range of hosts including humans
and a broad variety of animals (Tables 9.1 through 9.3). Spores, resistant infective form of E. bieneusi,
are ubiquitous in the environment. Consequently, humans can acquire infections of anthroponotic or
zoonotic origin through several transmission routes (person to person, animal to person, waterborne,
and foodborne). E. bieneusi has been identified in water sources as well as in wild, domestic, and
Biology of Foodborne Parasites
E. bieneusi Genotypes Identified Only in Humans
Genotype Name (GenBank
Accession Number) [Synonyms]
B (AF101198) [Type I]
Q (AF267147)a
R (AY945808)
S (AY945809)
T (AY945810)
U (AY945811)
V (AY945812)
W (AY945813)
Peru3 (AY371278)
Peru13 (EF014429)
Peru15 (EF014431)
CAF2 (DQ683747)
CAF3 (DQ683748)
CAF4 (DQ683749)
TypeIII (AY242477)
TypeV (AF242479)
NIA1 (EF458628)
HAN1 (EF458627)
UG2145 (AF502396)
Genotype 17 (EU140500)
S1 (FJ439677)
S2 (FJ439678)
S3 (FJ439680)
S4 (FJ439679)
S5 (FJ439681)
S7 (FJ439683)
S8 (FJ439684)
S9 (FJ439685)
CZ1 (GU198949)
CZ2 (GU198950)
CHN2 (HM992510)
Nig1 (JN997477)
Nig2 (JN997478)
Nig3 (JN997479)
Nig4 (JN997480)
Nig5 (JN997481)
MAY1 (JN595887)
KIN1 (JQ437573)
KIN2 (JQ437574)
KIN3 (JQ437575)
IH (KC708073)
Henan-II (JF691565)
SH1 (JX994257)
SH2 (JX994257)b
SH3 (JX994257)
Geographic Distribution
Australia, Cameroon, England, France, Germany,
the Netherlands, Nigeria, Switzerland, Tunisia
Germany, Switzerland
Gabon, Nigeria
Cameroon, Gabon
Brazil, Congo, Niger
Malawi, Uganda
the Netherlands
the Netherlands
the Netherlands
Czech Republic
Czech Republic
(Continued )
Enterocytozoon bieneusi
TABLE 9.1 (Continued )
E. bieneusi Genotypes Identified Only in Humans
Genotype Name (GenBank
Accession Number) [Synonyms]
SH4 (JX994257)
SH6 (JX994257)
SH8 (JX994257)
SH9 (JX994257)
SH10 (JX994257)
SH11 (JX994257)
SH12 (JX994257)
Geographic Distribution
ITS nucleotide sequence is 245 bp.
Incomplete ITS sequence.
food-producing farm animals, raising concerns of waterborne, foodborne, and zoonotic transmission.
Although, our understanding of the epidemiology of E. bieneusi is still limited, potential sources of
infection are beginning to be identified using molecular epidemiology for both humans and a wide range
of animals by comparison of genotypes from different hosts.17,40 The genotyping of E. bieneusi is a valuable tool for epidemiological investigations; it has shown that some E. bieneusi genotypes are associated
only with humans, while others infect both human and animals supporting the person-to-person and
animal-to-person transmission.17,40 A study of two different populations in Gabon (HIV-positive urban
population) and Cameroon (nonselected rural population) suggested that different genotypes may present different risks of infection associated with immune status and living conditions.6 Similar conclusions
were reached in a study that included samples from Niger and Vietnam in which it was suggested that
E. bieneusi genotype distribution in the two geographic areas was related to different routes of infection transmission, person to person in Niger and zoonotic in Vietnam.7 Evidence of a person-to-person
transmission was suggested in a study in a Thai orphanage because only genotype A was detected in all
positive samples.41
Current typing of E. bieneusi relies on molecular methods. Most of the genotyping studies performed
until now have been based on the analysis of the ribosomal internal transcriber spacer (ITS) nucleotide
sequences. The ITS is a fast-evolving genome region that has shown a high degree of diversity among
isolates.42 ITS data have allowed the identification of the host-adapted E. bieneusi genotypes in several
mammals, and other E. bieneusi genotypes with no host specificity, which are considered zoonotic. To
date, 204 E. bieneusi genotypes have been identified based on nucleotide sequence polymorphisms in
the 243 bp ITS region, including 52 in humans, 34 in both human and animals, and 106 host-adapted
genotypes in specific animal groups (Tables 9.1 through 9.3). Additionally, several genotypes have been
identified in water for which the host is still unknown (Table 9.4).
Numerous new genotypes have been named in the last 5 years. In 2008, 81 genotypes were recognized, the number increased to 93 genotypes by 2009, and at present they are 204 genotypes
(Tables 9.1 through 9.4).17,42 Caution should be taken when naming a new E. bieneusi genotype to
avoid confusions.42 The varying terminologies that have been used in the past to designate genotypes
make in many cases the comparison of published studies confusing and identification of new isolates
difficult. This lack of an acceptable nomenclature for naming ITS sequences resulted in multiple genotype designations being applied to the same ITS nucleotide sequence with some genotypes received
as many as six or seven different names (see genotypes D and TypeIV in Table 9.2). At the 10th
International Workshop on Opportunistic Pathogens, a roundtable discussion entitled “Terminology
for Enterocytozoon bieneusi” was held with the aim of proposing a plan to reduce confusion associated with the identification E. bieneusi genotypes. It was decided and published later in a consensus
report when multiple names appear for the same ITS sequence; the first recorded name for that
sequence will be the primary name with subsequent names listed as synonyms.42 For genotyping
purposes based on ITS sequence data, only the 243 bp of the ITS region should be used, and base
pairs associated with the small or large subunit rRNA should be excluded. It was also recommended
Biology of Foodborne Parasites
E. bieneusi Genotypes Identified in Humans and Animals
Genotype Name (GenBank
Accession Number)
A (AF101197)
Peru11 (AY371286)
Peru7 (AY371282)
CAF1 (DQ683746)
EbpC (AF076042)
E, Peru4,
WL13, WL17
Rhesus macaque
Meadow vole
Eastern cottontail
Peru16 (EF014427)
Peru10 (AY371285)
TypeIV (AF242478)
BEB-var, K,
Peru2, PtEBIII
Wild boar
Rhesus macaque
Guinea pig
Rhesus macaque
Cameroon, Gabon,
Germany, India, the
Netherlands, Niger,
Nigeria, Portugal, Peru,
Slovak Republic,
Switzerland, Thailand
Czech Republic
China, Peru, Thailand
United States
United States
United States
Gabon, Niger
Gabon, Korea
China, Peru, Thailand,
China, Germany, Japan,
Peru, Switzerland,
Austria, Poland
United States
United States
United States
United States
United States
Cameroon, China, France,
Gabon, England, Iran,
Malawi, the Netherlands,
Niger, Nigeria, Peru,
Portugal, Uganda
Korea, Portugal, United
Colombia, Germany,
Japan, Portugal
(Continued )
Enterocytozoon bieneusi
TABLE 9.2 (Continued )
E. bieneusi Genotypes Identified in Humans and Animals
Genotype Name (GenBank
Accession Number)
WL11 (AY237219)
O (AF267145)
PigEBITS7 (AF348475)
Peru6 (AY371281)
WL15 (AY237223)
D (AF101200)
CEbC, Peru9,
Meadow vole
Black bear
United States
United States
United States
United States
United States
United States
China, Germany,
China, India, Thailand
United States
Portugal, Peru
United States
Czech Republic
United States
United States
United States
United States
Brazil, Cameroon, China,
Congo, England, Gabon,
India, Iran, Malawi, the
Netherlands, Niger,
Nigeria, Portugal, Peru,
Poland, Russia, Spain,
Thailand, Tunisia,
China, Czech Republic,
Japan, United States
Czech Republic, Slovak
Argentina, Korea, South
United States
United States
United States
United States
River otter
United States
Rhesus macaque
Rhesus monkey
Wild boar
(Continued )
Biology of Foodborne Parasites
TABLE 9.2 (Continued )
E. bieneusi Genotypes Identified in Humans and Animals
Genotype Name (GenBank
Accession Number)
Rhesus macaque
S6 (FJ439682)
Peru8 (AY371283)
Rhesus macaque
CZ3 (GU198951)
C (AF101199)
BEB4 (AY331008)
EbpA (AF076040)
Wild boar
Abu Dhabi
Colombia, Czech
Czech Republic,
China, United States
Germany, Czech
China, Malawi, Nigeria,
Peru, Tunisia
Czech Republic, Germany
Czech Republic
Czech Republic,
France, Germany, the
Netherlands, Portugal,
Czech Republic,
China, Czech Republic
Argentina, China, South
Africa, United States
China, Czech Republic,
China, Czech Republic,
Germany, Japan,
Switzerland, United
Czech Republic, Poland
Czech Republic
Czech Republic,
Brazil, Czech Republic
(Continued )
Enterocytozoon bieneusi
TABLE 9.2 (Continued )
E. bieneusi Genotypes Identified in Humans and Animals
Genotype Name (GenBank
Accession Number)
PigITS5 (AF348173)
I (AF135836)
J (AF135837)
CHN3 (HM992511)
CHN4 (HM992512)
WL12 (AY237220)
PtEbII (DQ425108)
EbpD (AF076043)
WL7 (AY237215)
LW1 (JX000571)
Henan-I, SH7
Henan-III (JF691566)
Henan-IV (JQ029727)
Henan-V (JQ029728)
BEB6 (EU153584)
Rhesus macaque
Wild boar
Rhesus macaque
Rhesus macaque
Czech Republic
Japan, Korea, United
Czech Republic,
Argentina, China, Czech
Republic, Germany,
Korea, South Africa,
United States
Argentina, China,
Germany, Korea,
Portugal, United States
United States
United States
China, Switzerland
United States
United States
Biology of Foodborne Parasites
E. bieneusi Genotypes Identified Only in Animals
Genotype Name (GenBank Accession
Number) [Synonyms]
L (AF267142)
EbfelA (AF118144)
PtEbIV (DQ885580)
PtEbVIII (DQ885584)
D-like (DQ836345)
BEB3 (AY331007)a
BEB3-like (JQ923448)
PtEbXI (DQ885587)
M (AF267143)
N (AF267144)
4948 FL-2 2004 (DQ154136)
CEbA (EF139195)
CEbD (EF139198)
CEbF (EF139194)
BEB7 (EU153585)
BEB8 (JQ044398)
BEB9 (JQ044399)
BEB10 (KJ675196)
PtEbIX (DQ885585)
CHN5 (HM992513)
CHN6 (HM992514)
Horse1 (GQ406053)
Horse2 (GQ406054)
Horse3 (JQ804971)
Horse4 (JQ804972)
Horse5 (JQ804973)
Horse6 (JQ804974)
Horse7 (JQ804975)
Horse8 (JQ804976)
Horse9 (JQ804977)
Horse10 (JQ804978)
Horse11 (JQ804979)
H (AF135835) [PEbC]
G (AF135834)
PigITS1 (AF348469)
PigITS2 (AF348470)
PigITS3 (AF348471) [PEbB]
PigITS4 (AF348472)
PigITS6 (AF348474)
Wild boar
Geographic Distribution
United States
South Africa
United States
United States
United States
United States
Colombia, Japan, Portugal,
Switzerland, United States
Colombia, Czech Republic
Colombia, Czech Republic
Czech Republic
Czech Republic
Czech Republic
Czech Republic
Czech Republic
Czech Republic
Czech Republic
Czech Republic
Czech Republic
China, Germany, Japan,
Korea, Thailand
Czech Republic, Germany
Czech Republic
Czech Republic
United States
United States
Korea, United States
Korea, United States
United States
(Continued )
Enterocytozoon bieneusi
TABLE 9.3 (Continued )
E. bieneusi Genotypes Identified Only in Animals
Genotype Name (GenBank Accession
Number) [Synonyms]
PigITS8 (AF348476)
EbpB (AF076041)
CHN7 (HN992516)
CHN8 (HN992517)
CHN9 (HN992518)
CHN10 (HN992519)
E1 (EU849129)
F1 (EU883783)
CS-1 (KF607047)
CS-2 (KF607048)
CS-3 (KF607049)
CS-4 (KF607050)
CS-5 (KF607051)
CS-6 (KF607052)
CS-7 (KF607053)
CS-8 (KF607054)
HLJ-I (KJ475402)
HLJ-II (KJ475403)
HLJ-III (KJ475404)
HLJ-IV (KJ475401)
Wildboar1 (KF383396)
Wildboar2 (KF383397)
Wildboar3 (KF383398)
Wildboar4 (KF383400)
Wildboar5 (KF383402)
Wildboar6 (KF383404)
WL1 (AY237209)
WL2 (AY237210)
WL3 (AY237211)
WL4 (AY237212) [WL5]
WL6 (AY237214)
Wild boar
Wild boar
Wild boar
Wild boar
Wild boar
Wild boar
River otter
Black bear
River otter
Deer mouse
Eastern cottontail
White-tailed deer
Geographic Distribution
United States
Slovak Republic
Slovak Republic, Poland
Slovak Republic
United States
United States
United States
United States
United States
United States
United States
United States
United States
United States
United States
United States
United States
United States
United States
United States
United States
United States
United States
(Continued )
Biology of Foodborne Parasites
TABLE 9.3 (Continued )
E. bieneusi Genotypes Identified Only in Animals
Genotype Name (GenBank Accession
Number) [Synonyms]
WL9 (AY237217)
WL10 (AY237218)
WL14 (AY237222)
WL18 (KF591680)
WL19 (KF591681)
WL20 (KF591682)
WL21 (KF591683)
WL22 (KF591684)
WL23 (KF591685)
WL24 (KF591686)
WL25 (KF591686)
WL26 (KF591687)
PtEbV (DQ885589)
PtEbXII (DQ885588)
P (AF267146)
Peru6-var (DQ425107)
Col01a (AY668952)
Col02a (AY668953)
KB-1 (JF681175)
KB-2 (JF681176)
KB-3 (JF681177)
KB-4 (JF681178)
KB-5 (JF681179)
KB-6 (JF681180)
Macaque1 (JX000572)
Macaque2 (JX000573)
Macaque3 (KC441073)
Macaque4 (KC441074)
CM1 (KF305581)
CM2 (KF305586)
CM3 (KF305589)
CM4 (KF543866)
CM5 (KF543867)
CM6 (KF543870)
CM7 (KF543871)
Incomplete ITS nucleotide sequence.
White-tailed deer
White-tailed deer
Boreal red-backed vole
Boreal red-backed vole
Meadow vole
Deer mouse
Deer mouse
Rhesus macaque
Rhesus macaque
Cynomolgus monkey
Cynomolgus monkey
Cynomolgus monkey
Rhesus macaque
White-headed langur
Cynomolgus monkey
White-headed langur
Cynomolgus monkey
Rhesus macaque
Golden snub-nosed
Rhesus macaque
Rhesus macaque
Rhesus macaque
Geographic Distribution
United States
United States
United States
United States
United States
United States
United States
United States
United States
United States
United States
United States
United States
United States
United States
United States
United States
Enterocytozoon bieneusi
E. bieneusi Genotypes Identified in Water
Genotype Name (GenBank
Accession Number)
Water Geographic Origin
Reported Hosts in the Literaturea
Peru6 (AY371281)
Peru11 (AY371286)
Peru8 (AY371283)
EbpA (AF076040)
EbpC (AF076042)
China, Tunisia
China, Tunisia
China, Tunisia
EbpD (AF076043)
TypeIV (AF242478)
China, Ireland, Tunisia
C (AF101199)
D (AF101200)
China, Spain
China, Spain, Tunisia
D-like (DQ836345)
BEB3 (AY331007)
BEB6 (EU153584)
WL1 (AY237209)
WL2 (AY237210)
WL4 (AY237212)
WL6 (AY237214)
China, Tunisia
China, Tunisia, United
United States
Humans, bird, cattle, dog
Humans, baboon, rhesus macaque
Human, chicken, mouse
Human, cattle, horse, mouse, pig
Human, beaver, fox, muskrat,
rhesus macaque, otter, pig,
Human, pig
Human, cat, cattle, dog, rhesus
Human, mouse
Human, beaver, baboon, cattle,
dog, falcon, fox, horse, mouse,
muskrat, pig, raccoon, rhesus
Cattle, goat
WL12 (AY237220)
WL14 (AY237222)
WL15 (AY237223)
China, Tunisia
PtEbIV (DQ885580)
PtEbIX (DQ885585)
PigEBITS7 (AF348475)
PigEBITS8 (AF348476)
LW1 (JX000571)
WW1 (JQ863269)
WW2 (JQ863270)
WW3 (JQ863271)
WW4 (JQ863272)
WW5 (JQ863273)
WW6 (JQ863274)
WW7 (JQ863275)
WW8 (JQ863276)
WW9 (JQ863277)
SW1 (KF591677)
SW2 (KF591678)
SW3 (KF591680)
Muskrat, raccoon, squirrel,
Human, beaver, otter
Human, beaver, fox, horse,
muskrat, rhesus macaque, raccoon
Human, pig
Human, pig, wild boar
For details on geographic origin of humans and animals see Tables 9.2 and 9.3.
? Unknown.
Biology of Foodborne Parasites
that although names for new isolates should be at the discretion of the authors, each new name should
be clearly associated with a GenBank number and that submitters should clearly indicate the host and
geographic location.
Using phylogenetic analysis of the ITS region, five main groups (major clusters), numbered 1–5, were
described using genotype PtEbIX as an out-group.43 Group 1 was the largest of all five groups, and genotypes within this group have been isolated from a wide diversity of hosts worldwide, including humans
and domestic and wild animals. Groups 2, 3, and 4 contained at that time sequences isolated only from
animals and mostly from one host species. However, recent reports have found humans as hosts for some
of the genotypes included in those groups. For example, genotype BEB4, thought to be host specific
for cattle, has been also reported in humans and pigs.16,44 The other E. bieneusi genotype isolated from
humans outside group 1 was CAF4,43 although recently, more E. bieneusi genotypes in humans have
been placed outside group 1.5,45 Also, the number of groups has increased from 5 to 7.46 Another phylogenetic analysis using the 179 sequences of the ITS available at the moment in GenBank revealed 4 groups
clearly divergent from each other (groups 1–4).47 Group 1 included 94% of the sequences incorporated in
the study with many of these genotypes found in both humans and animals. The high diversity observed
in this group was mainly due to single-nucleotide polymorphisms with occasional insertion or deletion
of single base pairs. Sequences that constituted groups 2–5 were obtained from different hosts from different continents.
The use of a single genetic marker may have limitations in identifying genotypes that may have
different biologic characteristics.42,47,48 Hence, it is recommended that observations made using the
ITS need to be substantiated by using other genetic markers.42 Recently, a multilocus sequence typing (MLST) tool for E. bieneusi was developed targeting three microsatellite (MS1, MS3, and MS7)
and one minisatellite (MS4) markers in addition to the ITS marker49 (Table 9.5). Zoonotic and hostadapted genotypes as determined by ITS analysis mostly in previous studies were used.9,40,50–53 The
data generated supported the existence of two large groups: one that includes zoonotic genotypes and
another that includes host-adapted genotypes. Shortly after, another study used the same MLST tool
on 72 specimens from acquired immunodeficiency syndrome (AIDS) patients in Lima54; all specimens
previously genotyped using the ITS as A, D, IV, EbpC, WL11, Peru7, Peru8, Peru10, and Peru11.9
Micro- and minisatellite markers harbored varying degrees of sequence polymorphism; 28, 7, 7, 12,
Primer Sequences of Microsatellite and Minisatellite Loci Used in Multilocus Sequence
Typing (MLST) of E. bieneusi
Targeted Repeat
(TAT)31, (TAG)11
Primers (F, Forward; R, Reverse)
Product Size (bp)
Source: Modified from Feng, Y. et al., Appl. Environ. Microbiol., 77, 4822, 2011.
Enterocytozoon bieneusi
and 7 genotypes were identified using markets MS1, MS3, MS4, MS7, and ITS, respectively. The results
also indicated that all markets were in complete linkage disequilibrium.54 Multilocus analysis by concatenating sequences of all 5 markets identified 39 multilocus genotypes for the 72 specimens.54 An
additional MLST study has included specimens from Peru, Nigeria, India, and Kenya and showed a
lack of geographic segregation in E. bieneusi populations.55 The analysis revealed the presence again
of two subpopulations of E. bieneusi that differ from each other in population structures (epidemic vs.
clonal) and transmission routes (anthroponotic vs. zoonotic). Additional studies are necessary to better understand E. bieneusi epidemiology and to assess the relationship among MLST genotypes, host
specificity, virulence, and risk factors.
9.5.1 E. bieneusi in Humans
E. bieneusi is an opportunistic pathogen in HIV patients and other immunosuppressed individuals.
Opportunistic infections are a concern as the number of immunosuppressed individuals continues to
increase worldwide; the HIV pandemic continues to spread with the majority of these new infections
occurring in developing countries where access to active therapy is limited. In the meantime, the number of immunosuppressed individuals continues to increase in developed countries as a result of more
aggressive immunosuppressive therapies for organ transplants and immunologic disorders. HIV, the
most common cause of immunodeficiency worldwide, affects the CD4+ cell count. Once the counts are
below 200 cells/µL, patients are at risk of an opportunistic infection such as microsporidiosis. A clear
association between the presence of microsporidian infection and HIV seropositivity has been demonstrated in the literature particularly in patients with AIDS.8,56–58 In developed countries, prevalence of
E. bieneusi in HIV-seropositive patients is progressively decreasing, probably due to the access to effective highly active antiretroviral therapy (HAART).59 HAART results in suppression of HIV virus replication and CD4+ cell count restoration, which delays the onset, or eliminates the risk, of opportunistic
infections, including microsporidiosis. In a study from Australia, the prevalence of intestinal microsporidiosis in HIV-infected patients decreased from 11% in 1995 to 0% in 2004.59 However, in developing countries with limited access to HAART, microsporidiosis remains common.57,60,61 Infection with
E. bieneusi does not only affect HIV/AIDS patients but also affect patients with an impaired immune
system such as organ and bone marrow transplant recipients.11,62–64 Immunosuppressive therapy prescribed in solid organ and bone marrow transplant recipients leads to a deficiency of cellular immunity.
Infections with E. bieneusi in transplant recipients have been seen from days to years after transplantation.64 Therefore, transplant recipients undergoing immunosuppressive therapy should be considered a
risk group for acquisition of microsporidiosis, and in the case of persistent diarrhea, microsporidiosis
should be considered.
The percentage of microsporidian-positive children is usually higher than the percentage of
adults,8,57,65 with the highest occurrence of E. bieneusi in children aged 2 years or younger.37,65
A higher prevalence of infection was reported in Thailand among children 12–23 months old.41 This
could be explained by the fact that children have a developing immune system and generally have a
poorer hygiene with potential for higher exposure to microsporidian infections. Children in that age
range (12–23 months old) are also more active and have more independent eating habits compared
with those in the first year of age.
The identification of risk factors associated with infection is critical to understand modes of transmission. However, data concerning risk factors for intestinal microsporidiosis in humans are sparse.
A case-control study conducted in France to determine risk factor showed that intestinal microsporidiosis in persons with HIV infection and <200/mm3 CD4+ cells was associated with male homosexuality
and swimming, suggesting fecal–oral transmission, including sexual and waterborne routes.56 Zoonotic
transmission has been suggested in many studies5,40,65 with at least one case of zoonotic E. bieneusi
infection reported.40 Zoonotic transmission was the most likely cause of infection of a 2-year-old child in
Peru based on the uniqueness of the genotype identified (Peru16) in multiple guinea pigs from unrelated
households as well as in guinea pigs in the household, the close contacts between the child and infected
guinea pigs, and the absence of genotype Peru16 in other children in the community.40 A study of HIV/
AIDS patients with intestinal microsporidiosis suggested that the possibility of zoonotic transmission of
Biology of Foodborne Parasites
E. bieneusi as household presence of and contact with droppings of domestic animals (especially ducks
and chickens) was found to be a risk factor for infection.50 A study showed that 90% of E. bieneusi infections in farmers were caused by genotypes commonly found in animals, including all three dominant
genotypes (EbpC, D, and TypeIV), PigEBITS7, and EbpD.10 In China, contact with pigs was strongly
associated with the most prevalent genotype, EbpC, which is a common genotype in pigs.10 However, in
studies conducted in Thailand, Nigeria, and France, contact with animals was not identified as a risk factor for E. bieneusi infection.5,39,56 An association between infection and poor sanitary conditions (lack of
a flush toilet, municipal garbage collection, and running water) has also been suggested.50 Hand washing
before meals was a protective factor against microsporidiosis.10
Diarrhea is the most common clinical symptom associated with E. bieneusi infections, but several
studies suggest that E. bieneusi infection can also remain asymptomatic.8,13,16,38,66 Therefore, the role
of asymptomatic carriers as a source of infection is important and maybe underrated. The reactivation
of latent infection in asymptomatic chronic carriers due to an immunosuppression is also a possibility. Diagnosis of microsporidiosis when a single sample is collected could be challenging because
spore production is not a constant process as demonstrated in asymptomatic children followed for
2 months.38
9.5.2 E. bieneusi in Animals
Numerous animal hosts have been identified for E. bieneusi, including domestic and wild animals.17
Comparison of prevalence described in the literature should be treated with caution because different
studies used different populations, different specimens, and different diagnostic tests. Tables 9.2 and 9.3
include all genotypes that have been identified in animals including information about the countries in
which they have been reported. Some genotypes are associated with specific groups of animals indicating that cross transmission of those genotypes among different groups of animals is probably limited. However, other genotypes are not host specific with some of them able to infect both animals and
humans. Therefore, the role of animals in the transmission of human infections requires further study. Nonhuman Primates
The first report of infection with E. bieneusi in nonhuman primates was described in two simian immunodeficiency virus (SIV)–infected rhesus macaques experimentally infected with E. bieneusi spores
from an AIDS patient.67 Both animals began shedding spores within a week of inoculation proving the
possibility of using SIV-infected rhesus monkey as an animal model to study E. bieneusi infection in
HIV/AIDS patients. Soon after this report, the first natural infection of this parasite in nonhuman primates was documented in 18 SIV-infected macaques with hepatobiliary disease.68 It was also found as
the cause of proliferative serositis in seven SIV-infected macaques with peritonitis.69 In a recent study in
free-range rhesus monkeys in a public park in China, E. bieneusi was detected in 28.2% of fecal specimens examined indicating these animals might be reservoirs for humans.70 Also in China, 1386 fecal
specimens of nonhuman primates were examined, and E. bieneusi was detected in 158 (11.4%) specimens of five nonhuman primate species.71 E. bieneusi was also found in 12.3% of 235 newly captured
baboons in Kenya.72
Thirty genotypes have been reported in nonhuman primates. Eighteen of them appear to be host specific to nonhuman primates (Table 9.3), while the other 12 genotypes (A, D, I, BEB6, EbpC, Henan-V,
Peru7, Peru8, Peru11, TypeIV, PigEBITS7, and WL15) have also been reported in humans implying that
nonhuman primates might be reservoirs for human microsporidiosis. Companion Animals
There are few published reports of microsporidia in companion animals. In cats, prevalence of E. bieneusi
ranges from 5% to 17% in studies conducted in Colombia, Germany, Japan, and Switzerland.73–76
Additionally, E. bieneusi has been also reported in six cats in Portugal.53 In dogs, prevalence of E. bieneusi
Enterocytozoon bieneusi
ranges from 2.5% to 15% in studies conducted in Colombia, Japan, Spain, and Switzerland.74,75,77–79 In
horses, the first two studies that examined fecal specimens from Switzerland and Spain found all animals
negative.78,80 However, later studies reported a prevalence of 10.8% and 17.3% in horses from Colombia
and the Czech Republic, respectively.81,82
Because cats, dogs, and horses are parasitized by both host-specific and zoonotic E. bieneusi genotypes (Tables 9.2 and 9.3), the role of domestic animals in the transmission of human infections requires
further study. Properly designed studies are needed to assess the role of companion animals as a source
of zoonotic genotypes of E. bieneusi. Livestock
E. bieneusi was first detected in eight calves in Germany.76,83 Since then, it is clear that E. bieneusi is
a common parasite of dairy and beef cattle.84,85 It has been detected in cattle from countries in Africa,
Asia, Europe, North America, and South America (Tables 9.2 and 9.3). Prevalence of E. bieneusi in
cattle has a wide range from 14.3% in Argentina86 to 37.6% in China.44 Prevalence appears to vary also
among age groups. Prevalence was found higher in postweaned calves (13%) and heifers (23%) than in
preweaned calves (3%) and adults (4.4%).87–90 In a longitudinal study of 30 cattle, healthy animals were
examined at weekly, bimonthly, and then monthly intervals from 1 week to 24 months of age for the
presence of E. bieneusi. The overall prevalence was 24% with lower prevalence in preweaned calves
(11.7%) and highest in postweaned calves (44.4%). However, the cumulative prevalence of E. bieneusi
in those animals was 100% since spores of E. bieneusi were detected in all 30 calves at some time during the study.
The presence of E. bieneusi in milk has been documented in Korea,91 giving rise to the concern that
dairy cow’s milk may play a significant role in the transmission of E. bieneusi infections to humans.
The extent to which E. bieneusi–infected cattle might pose a risk of human infection is still unknown.
Genotypes identified in cattle are mostly cattle-specific genotypes (Table 9.3), but genotypes identified in
humans, such as TypeIV and D, have also been identified in various studies in cattle (Table 9.2). Besides,
genotypes that were thought to have only cattle as a host such as I, J, BEB417 have now also been reported
in humans as well as other animals (Table 9.2).
Information about the infection with E. bieneusi in small ruminants is scarce. There are only two
reports of E. bieneusi in two goats in Spain78 and in Peru49 and none in sheep.
In swine, E. bieneusi was first detected in four immunocompetent pigs in Switzerland.92 Also in
Switzerland, E. bieneusi was detected in 34.8% of 109 fecal samples examined from healthy pigs.80 In
the United States, feces and bile from pigs were examined for the presence of E. bieneusi in a slaughterhouse.93 Sixty-four of the 202 (31.6%) pigs were found positive, and the viability of these spores was
demonstrated by their capability to infect gnotobiotic piglets. A higher prevalence was found in pigs in
the Czech Republic with 94% of them positive for E. bieneusi by PCR.94 The presence of E. bieneusi has
been recognized in pigs in all age categories, and it is likely that piglets may acquire the infection from
their mothers at a very early age.94 Recently, it was reported that infection rates in nursery pigs (63.6%)
were significantly higher than in preweaned (41%) and growing (26.3%) pigs.95 E. bieneusi spore excretion was monitored weekly for 2 months in three naturally infected pigs; spores were detected in 67% of
samples showing that shedding of spores was intermittent.80 Significant higher prevalence of E. bieneusi
was found in piglets aged 2–3.9 months than in pigs of other age groups.39 Most animals appeared to
be in good health condition with no association between occurrence of diarrhea and E. bieneusi infections.93,94 In Korea, the parasite was identified in 16% of the 472 piglet without diarrhea and in 12% of
the 235 piglets with diarrhea; however, it was unclear if E. bieneusi was the cause of diarrhea because
other pathogens were not examined.96
Numerous genotypes of E. bieneusi have been reported in pigs (Tables 9.2 and 9.3), some with potential to be zoonotic and others are apparently host specific. A study in a farm community in Thailand that
included sampling of pigs and humans did not identified zoonotic transmission.39 However, infection in
humans with zoonotic genotype EbpC was associated with having pigs in the household.10 Clearly, more
studies are necessary to evaluate the role of pigs as a source of infection for humans.
Biology of Foodborne Parasites Birds
The first case of E. bieneusi in birds was detected in underweight broiler chickens.97 This constituted
the first detection of this parasite in nonmammalian hosts. Since then, it has been identified in captive
exotic birds of the orders Passeriformes and Psittaciformes in Brazil, Czech Republic, and Portugal98–100;
pigeons in the Netherlands, Brazil, Iran, Portugal, Spain, and the United States99–104; and falcons in the
United Arab Emirates.105 Most infected birds were apparently healthy and might serve as asymptomatic
carriers and a source of environmental contamination. Additionally, the finding of genotypes in birds,
previously identified in humans, such as A, Peru6, or EbpA, suggests that there could be a possible
transmission of E. bieneusi between birds and humans (Table 9.2). A connection between contact with
ducks and chickens and infection with genotype Peru6 was found during an investigation of HIV-positive
patients from two hospitals in Peru.50 Other Animals
There are sporadic reports on the presence of E. bieneusi in other wild and domestic animals. E. bieneusi
was detected in the feces of fur-bearing mammals, including raccoons, muskrats, beavers, foxes, and
otters, from Maryland, United States.52 The study found mostly host-adapted genotypes but also genotypes previously described in humans and other animals. A recent study detected E. bieneusi in 29% of
the 255 fecal specimens examined from wildlife living in the watershed of the New York City water supply system.106 These animals belonged to 23 species in 6 orders; most of the positive animals (64/74) were
rodents and carnivores, while only 10 were other mammals. The dominant E. bieneusi genotype was
WL4 that was also the dominant genotype found in stormwater samples in the same study, suggesting
that wildlife played a significant role in contamination of water by E. bieneusi in the watershed. Because
wild mammals live in or near surface water, they can be a source of contamination in water with human
pathogenic E. bieneusi.52,106 In Peru, 59 guinea pigs in 20 households were examined; 3 guinea pigs
from 2 unrelated households carried Peru16 suggesting that guinea pigs are the natural host of Peru16.40
This unusual genotype was also found in a child and seven guinea pigs in the same household, strongly
suggesting the possibility of zoonotic transmission.40 Wild and domestic rabbits were found positive
for E. bieneusi at four different geographic locations in Spain.79 Eurasian wild boars from four central
European countries were found to be E. bieneusi positive.107 A host-specific genotype, P, was identified
in a llama from Germany.76
9.5.3 E. bieneusi in Water
Spores of microsporidia have been detected in surface water, groundwater, and tertiary agricultural
effluent,108–111 thus posing a contamination risk to drinking, recreational, and agricultural water supplies.
There is one report of a presumed waterborne outbreak with microsporidia that affected 200 persons in
which the major factor associated with microsporidiosis infection was living in an area serviced by 1
of the 3 distribution subsystems in town.20 Data on molecular characterization of E. bieneusi in water
samples are presented in Table 9.4. In Spain, E. bieneusi was found in 16.5% of the 109 water samples
from drinking water plants, wastewater treatment plants, and locations of influence on 4 river basins
examined during a year-longitudinal study.112 Finished waters from drinking water plants showed a lower
prevalence of microsporidia. In the study, they observed a tendency of microsporidia to raise in spring
and summer with higher prevalence in both raw and finished waters from wastewater treatment plants.
In China, E. bieneusi was detected in more than 90% of wastewater samples in Shanghai, Qingdao, and
Wuhan.46 The majority of E. bieneusi found in this study belonged to group 1; within this group, the most
prevalent genotype was D, which has a wide geographic distribution and host range that includes humans
and wild and domestic animals. In Tunisia, a survey in wastewater detected E. bieneusi in 62% of 232
samples examined that included raw water, treated wastewater, dry sludge, and dehydrated sludge.113 The
presence of E. bieneusi in surface water suggests the possibility that domestic and wild animals could
be sources of contamination. E. bieneusi was detected in 13 of 23 water samples collected from a lake
where monkeys bathed; all genotypes found in the water samples were also identified in the monkeys.70
Enterocytozoon bieneusi
Irrigation water may become contaminated by sewage or runoff from numerous sources. Likewise,
rain events may carry fecal contamination from farms and domestic and wild animals into sources of
irrigation water. A study on irrigation water from the United States and several countries in Central
America used for crop production demonstrated that 28% of the investigated waters tested positive for
microsporidia.109 Crops may come in contact with microsporidian spores if irrigation water is contaminated. If those fruits and vegetables need minimal process or are eaten raw, there is a risk of disease to
consumers who consume these products.
Accidental ingestion of water while bathing carries a risk of infection by waterborne pathogens.
However, limited information on the presence of microsporidia in recreational water is available.
In pools, microsporidia spores most likely can escape malfunctioned filtration systems because they
are very small and the effects of routine disinfection on microsporidian spore viability are unknown.
A recent study showed that chlorine is effective in killing spores of E. intestinalis.114 A 1-year prospective study in swimming pools in France showed a low level of contamination with microsporidia.115
However, another study showed that intestinal microsporidiosis was associated with swimming in
pools, suggesting the likely occurrence of fecal–oral transmission through contaminated water.56
Swimming in freshwater is also a potential risk of acquiring waterborne pathogens. A 1-year prospective study at two recreational lakes and three river sites located near Paris, where bathing and boating
are frequent, showed the presence of E. bieneusi in only one lake sample and one river sample, suggesting that the risk of human infection by microsporidia during swimming at these sites was low.110
However, the risk of microsporidian infection by swimming in either freshwater or swimming pools
needs further evaluation.
9.5.4 E. bieneusi in Food
Microsporidia has been found in fresh produce sold at the retail level in Poland, including grocery stores,
street vendors, market stalls, and supermarkets. Specifically, spores of E. bieneusi were identified in
raspberries and curly lettuce sold at a market stall and in mung bean sprouts from a supermarket.18 Only
one outbreak of microsporidiosis associated with foodborne transmission has been reported in the literature.19 The outbreak affected persons visiting a hotel in Sweden, and E. bieneusi was the only pathogen
detected in stools despite extensive testing. In all samples in which sequencing was conducted, genotype
C was identified. Of the individual food items, cheese sandwich served during morning coffee break
and salad served during lunch had the highest relative risks. Both cheese sandwich and salad contained
cucumber slices that could be the probable vehicle of transmission.
It is possible that E. bieneusi may be an underreported cause of gastrointestinal outbreaks, given that
for over half of the reported foodborne outbreaks, the etiological agent is unknown. The long incubation
period of E. bieneusi (3–15 days) delays the recognition of outbreak occurrence. As a consequence of the
delay, it is harder to acquire information relevant for an epidemiological investigation and to have access
to suspected food items for testing.
9.6 Pathogenesis and Clinical Features
E. bieneusi infections are mostly limited to the gastrointestinal tract. The most common primary sites
of infection are epithelial cells lining the duodenum and jejunum. Diarrhea is the main clinical symptom associated with E. bieneusi infections, although the mechanism by which it induces diarrhea has
not been determined. Most likely, the pathogenesis of the intestinal disease is related to apoptosis of
enterocytes as a result of cellular infection. Besides infecting the gastrointestinal tract, E. bieneusi can
occasionally affect the respiratory tract causing pulmonary infection. All documented cases of pulmonary microsporidiosis were reported in immunocompromised patients.116,117 Main clinical symptoms
are persistent cough, nonpurulent sputum, dyspnea, and wheezing.118 However, symptomatology is not
always associated with respiratory infection.117
E. bieneusi is recognized as an opportunistic pathogen in patients with AIDS and in other immunocompromised individuals such as organ transplant recipients.5,11,57,60,64 Patients with low CD4 cell counts
Biology of Foodborne Parasites
are most susceptible to intestinal microsporidiosis.5,7,50,56 In those patients, infection with E. bieneusi
can be serious and life threatening. The clinical features associated with E. bieneusi infection include
persistent diarrhea, abdominal pain, fever, and weight loss.5,119 In HIV-positive children with E. bieneusi,
wasting was twice more likely than in children without infection.120 Also, infection of the biliary tract
has been reported as the cause of cholangiopathy including sclerosing cholangitis, papillary stenosis,
cholecystitis, and cholangitis.121,122
Immunocompetent individuals usually develop a self-limiting disease and may have diarrhea that usually resolve within a few weeks or months. Several studies have also reported no significant association
between the presence of E. bieneusi in fecal specimens and diarrhea,8,66 suggesting that infection can be
asymptomatic. However, it is still unknown whether certain genotypes are pathogenic, whereas others
are not.
9.7 Treatment and Prevention
The treatment of E. bieneusi infection is still a problem. Clinical microsporidiosis has mostly been
reported in immunocompromised patients such as patients with AIDS or organ transplant recipients. For
those patients, the most important treatment is the restoration of immune functions. Cellular immune
responses have been considered essential for the control and elimination of E. bieneusi in patients with
HIV.123 Since the introduction of HAART, there has been a declining incidence of intestinal microsporidiosis and reduction in AIDS-related mortality because of restoration of cell-mediated immunity.59
In transplant recipients, a suspension of immunosuppressive treatment may be necessary in controlling
Albendazole, which is effective against microsporidia other than E. bieneusi, seems to alleviate diarrhea in E. bieneusi–infected patients without clearing the infection, but decreasing the number of spores
in feces.125 Albendazole therapy brought symptomatic relief but no microbiological clearance to a liver
transplant recipient with chronic diarrhea associated with E. bieneusi infection.126 Fumagillin is an antibiotic originally used against Nosema apis infections in honeybees. Fumagillin (20 mg three times a day)
has been an effective treatment for chronic diarrhea due to E. bieneusi infection in immunocompromised
patients.12,127 However, efficacy of fumagillin is counterbalance by its adverse effects as fumagillin treatment has been associated to hematological toxicity.12,64 Patients with preexisting cytopenia require close
thrombocytes monitoring. In addition, fumagillin was suspected to be the cause of a case of meningoencephalitis in a renal transplant recipient with E. bieneusi infection.128 Further research is needed to
establish proper treatment protocols and manage side effects of these treatments.
In persons at risk for developing life-threatening microsporidiosis, such as those with AIDS or organ
transplant recipients, prevention should be established in terms of hygiene to avoid oral–fecal transmission such as hand washing, washing fresh vegetables, drinking boiled or bottled water, and limiting
contact with animals susceptible to transmission of E. bieneusi.
1. Mathis, A., Weber, R., and Deplazes, P. Zoonotic potential of the microsporidia. Clin Microbiol Rev 18,
423–445 (2005).
2. Didier, E.S. and Weiss, L.M. Microsporidiosis: Current status. Curr Opin Infect Dis 19, 485–492 (2006).
3. Molina, J.M. et al. Intestinal microsporidiosis in human immunodeficiency virus-infected patients with
chronic unexplained diarrhea: Prevalence and clinical and biologic features. J Infect Dis 167, 217–221
4. Van Gool, T. et al. High prevalence of Enterocytozoon bieneusi infections among HIV-positive individuals with persistent diarrhoea in Harare, Zimbabwe. Trans R Soc Trop Med Hyg 89, 478–480 (1995).
5. Akinbo, F.O. et al. Molecular epidemiologic characterization of Enterocytozoon bieneusi in HIV-infected
persons in Benin City, Nigeria. Am J Trop Med Hyg 86, 441–445 (2012).
6. Breton, J. et al. New highly divergent rRNA sequence among biodiverse genotypes of Enterocytozoon
bieneusi strains isolated from humans in Gabon and Cameroon. J Clin Microbiol 45, 2580–2589 (2007).
Enterocytozoon bieneusi
7. Espern, A. et al. Molecular study of microsporidiosis due to Enterocytozoon bieneusi and Encephalitozoon
intestinalis among human immunodeficiency virus-infected patients from two geographical areas:
Niamey, Niger, and Hanoi, Vietnam. J Clin Microbiol 45, 2999–3002 (2007).
8. Lobo, M.L., Xiao, L., Antunes, F., and Matos, O. Microsporidia as emerging pathogens and the implication for public health: A 10-year study on HIV-positive and -negative patients. Int J Parasitol 42,
197–205 (2012).
9. Sulaiman, I.M. et al. A molecular biologic study of Enterocytozoon bieneusi in HIV-infected patients in
Lima, Peru. J Eukaryot Microbiol 50(Suppl.), 591–596 (2003).
10. Wang, L. et al. Zoonotic Cryptosporidium species and Enterocytozoon bieneusi genotypes in HIVpositive patients on antiretroviral therapy. J Clin Microbiol 51, 557–563 (2013).
11. Galvan, A.L. et al. First cases of microsporidiosis in transplant recipients in Spain and review of the
literature. J Clin Microbiol 49, 1301–1306 (2011).
12. Lanternier, F. et al. Microsporidiosis in solid organ transplant recipients: Two Enterocytozoon bieneusi
cases and review. Transpl Infect Dis 11, 83–88 (2009).
13. Nkinin, S.W., Asonganyi, T., Didier, E.S., and Kaneshiro, E.S. Microsporidian infection is prevalent in
healthy people in Cameroon. J Clin Microbiol 45, 2841–2846 (2007).
14. Desportes, I. et al. Occurrence of a new microsporidian: Enterocytozoon bieneusi n. g., n. sp., in the
enterocytes of a human patient with AIDS. J Protozool 32, 250–254 (1985).
15. Albrecht, H. and Sobottka, I. Enterocytozoon bieneusi infection in patients who are not infected with
human immunodeficiency virus. Clin Infect Dis 25, 344 (1997).
16. Sak, B. et al. Unapparent microsporidial infection among immunocompetent humans in the Czech
Republic. J Clin Microbiol 49, 1064–1070 (2011).
17. Santin, M. and Fayer, R. Microsporidiosis: Enterocytozoon bieneusi in domesticated and wild animals.
Res Vet Sci 90, 363–371 (2011).
18. Jedrzejewski, S., Graczyk, T.K., Slodkowicz-Kowalska, A., Tamang, L., and Majewska, A.C. Quantitative
assessment of contamination of fresh food produce of various retail types by human-virulent microsporidian spores. Appl Environ Microbiol 73, 4071–4073 (2007).
19. Decraene, V., Lebbad, M., Botero-Kleiven, S., Gustavsson, A.M., and Lofdahl, M. First reported foodborne outbreak associated with microsporidia, Sweden, October 2009. Epidemiol Infect 140, 519–527
20. Cotte, L. et al. Waterborne outbreak of intestinal microsporidiosis in persons with and without human
immunodeficiency virus infection. J Infect Dis 180, 2003–2008 (1999).
21. Wasson, K. and Peper, R.L. Mammalian microsporidiosis. Vet Pathol 37, 113–128 (2000).
22. Keeling, P.J. and Fast, N.M. Microsporidia: Biology and evolution of highly reduced intracellular parasites. Ann Rev Microbiol 56, 93–116 (2002).
23. Rijpstra, A.C., Canning, E.U., Van Ketel, R.J., Eeftinck Schattenkerk, J.K., and Laarman, J.J. Use of
light microscopy to diagnose small-intestinal microsporidiosis in patients with AIDS. J Infect Dis 157,
827–831 (1988).
24. Vossbrinck, C.R., Maddox, J.V., Friedman, S., Debrunner-Vossbrinck, B.A., and Woese, C.R. Ribosomal
RNA sequence suggests microsporidia are extremely ancient eukaryotes. Nature 326, 411–414 (1987).
25. Gill, E.E. and Fast, N.M. Assessing the microsporidia-fungi relationship: Combined phylogenetic analysis of eight genes. Gene 375, 103–109 (2006).
26. Fischer, W.M. and Palmer, J.D. Evidence from small-subunit ribosomal RNA sequences for a fungal
origin of Microsporidia. Mol Phylogenet Evol 36, 606–622 (2005).
27. Capella-Gutierrez, S., Marcet-Houben, M., and Gabaldon, T. Phylogenomics supports microsporidia as
the earliest diverging clade of sequenced fungi. BMC Biol 10, 47 (2012).
28. Modigliani, R. et al. Diarrhoea and malabsorption in acquired immune deficiency syndrome: A study of
four cases with special emphasis on opportunistic protozoan infestations. Gut 26, 179–187 (1985).
29. Canning, E. and Hollister, W. Royal society of tropical medicine and hygiene meeting at Manson House,
London, 16 March 1989 New intestinal protozoa—Coccidia and microsporidia. Trans R Soc Trop Med
Hyg 84, 181–186 (1990).
30. Akiyoshi, D.E. et al. Genomic survey of the non-cultivatable opportunistic human pathogen,
Enterocytozoon bieneusi. PLoS Pathog 5, e1000261 (2009).
31. Weber, R., Bryan, R.T., Schwartz, D.A., and Owen, R.L. Human microsporidial infections. Clin Microbiol
Rev 7, 426–461 (1994).
Biology of Foodborne Parasites
32. Sak, B. et al. Seropositivity for Enterocytozoon bieneusi, Czech Republic. Emerg Infect Dis 16, 335–337
33. Kucerova, Z. et al. Microsporidiosis and cryptosporidiosis in HIV/AIDS patients in St. Petersburg,
Russia: Serological identification of microsporidia and Cryptosporidium parvum in sera samples from
HIV/AIDS patients. AIDS Res Hum Retrovir 27, 13–15 (2011).
34. Ghosh, K. and Weiss, L.M. Molecular diagnostic tests for microsporidia. Interdisc Perspect Infect Dis
2009, 926521 (2009).
35. Carville, A. et al. Development and application of genetic probes for detection of Enterocytozoon
bieneusi in formalin-fixed stools and in intestinal biopsy specimens from infected patients. Clin Diagn
Lab Immunol 4, 405–408 (1997).
36. Rinder, H. et al. Blinded, externally controlled multicenter evaluation of light microscopy and PCR for
detection of microsporidia in stool specimens. The Diagnostic Multicenter Study Group on Microsporidia.
J Clin Microbiol 36, 1814–1818 (1998).
37. Pagornrat, W. et al. Carriage rate of Enterocytozoon bieneusi in an orphanage in Bangkok, Thailand.
J Clin Microbiol 47, 3739–3741 (2009).
38. Mungthin, M. et al. Spore shedding pattern of Enterocytozoon bieneusi in asymptomatic children. J Med
Microbiol 54, 473–476 (2005).
39. Leelayoova, S. et al. Genotypic characterization of Enterocytozoon bieneusi in specimens from pigs and
humans in a pig farm community in Central Thailand. J Clin Microbiol 47, 1572–1574 (2009).
40. Cama, V.A. et al. Transmission of Enterocytozoon bieneusi between a child and guinea pigs. J Clin
Microbiol 45, 2708–2710 (2007).
41. Leelayoova, S. et al. Transmission of Enterocytozoon bieneusi genotype a in a Thai orphanage. Am J Trop
Med Hyg 73, 104–107 (2005).
42. Santin, M. and Fayer, R. Enterocytozoon bieneusi genotype nomenclature based on the internal transcribed spacer sequence: A consensus. J Eukaryot Microbiol 56, 34–38 (2009).
43. Thellier, M. and Breton, J. Enterocytozoon bieneusi in human and animals, focus on laboratory identification and molecular epidemiology. Parasite (Paris, France) 15, 349–358 (2008).
44. Zhang, X. et al. Identification and genotyping of Enterocytozoon bieneusi in China. J Clin Microbiol 49,
2006–2008 (2011).
45. Pomares, C. et al. A new and highly divergent Enterocytozoon bieneusi genotype isolated from a renal
transplant recipient. J Clin Microbiol 50, 2176–2178 (2012).
46. Li, N. et al. Molecular surveillance of Cryptosporidium spp., Giardia duodenalis, and Enterocytozoon
bieneusi by genotyping and subtyping parasites in wastewater. PLoS Neglect Trop Dis 6, e1809 (2012).
47. Henriques-Gil, N., Haro, M., Izquierdo, F., Fenoy, S., and del Aguila, C. Phylogenetic approach to the
variability of the microsporidian Enterocytozoon bieneusi and its implications for inter- and intrahost
transmission. Appl Environ Microbiol 76, 3333–3342 (2010).
48. Widmer, G. and Akiyoshi, D.E. Host-specific segregation of ribosomal nucleotide sequence diversity in
the microsporidian Enterocytozoon bieneusi. Infect Genet Evol 10, 122–128 (2010).
49. Feng, Y. et al. Development of a multilocus sequence typing tool for high-resolution genotyping of
Enterocytozoon bieneusi. Appl Environ Microbiol 77, 4822–4828 (2011).
50. Bern, C. et al. The epidemiology of intestinal microsporidiosis in patients with HIV/AIDS in Lima, Peru.
J Infect Dis 191, 1658–1664 (2005).
51. Sulaiman, I.M. et al. Molecular characterization of Enterocytozoon bieneusi in cattle indicates that only
some isolates have zoonotic potential. Parasitol Res 92, 328–334 (2004).
52. Sulaiman, I.M. et al. Molecular characterization of microsporidia indicates that wild mammals harbor
host-adapted Enterocytozoon spp. as well as human-pathogenic Enterocytozoon bieneusi. Appl Environ
Microbiol 69, 4495–4501 (2003).
53. Lobo, M.L. et al. Genotypes of Enterocytozoon bieneusi in mammals in Portugal. J Eukaryot Microbiol
53(Suppl. 1), S61–64 (2006).
54. Li, W. et al. Population genetic analysis of Enterocytozoon bieneusi in humans. Int J Parasitol 42, 287–
293 (2012).
55. Li, W. et al. Multilocus sequence typing of Enterocytozoon bieneusi: Lack of geographic segregation and
existence of genetically isolated sub-populations. Infect Genet Evol 14, 111–119 (2013).
56. Hutin, Y.J. et al. Risk factors for intestinal microsporidiosis in patients with human immunodeficiency
virus infection: A case-control study. J Infect Dis 178, 904–907 (1998).
Enterocytozoon bieneusi
57. Tumwine, J.K. et al. Enterocytozoon bieneusi among children with diarrhea attending Mulago Hospital
in Uganda. Am J Trop Med Hyg 67, 299–303 (2002).
58. Saigal, K. et al. Intestinal microsporidiosis in India: A two year study. Parasitol Int 62, 53–56 (2013).
59. van Hal, S.J. et al. Declining incidence of intestinal microsporidiosis and reduction in AIDS-related
mortality following introduction of HAART in Sydney, Australia. Trans R Soc Trop Med Hyg 101, 1096–
1100 (2007).
60. Agholi, M., Hatam, G.R., and Motazedian, M.H. HIV/AIDS-associated opportunistic protozoal diarrhea.
AIDS Res Hum Retrovir 29, 35–41 (2013).
61. Wumba, R. et al. Epidemiology, clinical, immune, and molecular profiles of microsporidiosis and cryptosporidiosis among HIV/AIDS patients. Int J Gen Med 5, 603–611 (2012).
62. Rabodonirina, M. et al. Enterocytozoon bieneusi as a cause of chronic diarrhea in a heart-lung transplant
recipient who was seronegative for human immunodeficiency virus. Clin Infect Dis 23, 114–117 (1996).
63. Agholi, M., Hatam, G.R., and Motazedian, M.H. Microsporidia and coccidia as causes of persistence
diarrhea among liver transplant children: Incidence rate and species/genotypes. Pediatr Infect Dis J 32,
185–187 (2013).
64. Champion, L. et al. Fumagillin for treatment of intestinal microsporidiosis in renal transplant recipients.
Am J Transplant 10, 1925–1930 (2010).
65. Maikai, B.V. et al. Molecular characterizations of Cryptosporidium, Giardia, and Enterocytozoon in
humans in Kaduna State, Nigeria. Exp Parasitol 131, 452–456 (2012).
66. Sarfati, C. et al. Prevalence of intestinal parasites including microsporidia in human immunodeficiency
virus-infected adults in Cameroon: A cross-sectional study. Am J Trop Med Hyg 74, 162–164 (2006).
67. Tzipori, S. et al. Transmission and establishment of a persistent infection of Enterocytozoon bieneusi,
derived from a human with AIDS, in simian immunodeficiency virus-infected rhesus monkeys. J Infect
Dis 175, 1016–1020 (1997).
68. Mansfield, K.G. et al. Identification of an Enterocytozoon bieneusi-like microsporidian parasite in
simian-immunodeficiency-virus-inoculated macaques with hepatobiliary disease. Am J Pathol 150,
1395–1405 (1997).
69. Chalifoux, L.V. et al. Enterocytozoon bieneusi as a cause of proliferative serositis in simian immunodeficiency virus-infected immunodeficient macaques (Macaca mulatta). Arch Pathol Lab Med 124, 1480–
1484 (2000).
70. Ye, J. et al. Anthroponotic enteric parasites in monkeys in public park, China. Emerg Infect Dis 18,
1640–1643 (2012).
71. Karim, M.R. et al. Genetic polymorphism and zoonotic potential of Enterocytozoon bieneusi from nonhuman primates in China. Appl Environ Microbiol 80, 1893–1898 (2014).
72. Li, W. et al. Cyclospora papionis, Cryptosporidium hominis, and human-pathogenic Enterocytozoon
bieneusi in captive baboons in Kenya. J Clin Microbiol 49, 4326–4329 (2011).
73. Santin, M., Trout, J.M., Vecino, J.A., Dubey, J.P., and Fayer, R. Cryptosporidium, Giardia and
Enterocytozoon bieneusi in cats from Bogota (Colombia) and genotyping of isolates. Vet Parasitol 141,
334–339 (2006).
74. Abe, N., Kimata, I., and Iseki, M. Molecular evidence of Enterocytozoon bieneusi in Japan. J Vet Med
Sci/Japan Soc Vet Sci 71, 217–219 (2009).
75. Mathis, A., Breitenmoser, A.C., and Deplazes, P. Detection of new Enterocytozoon genotypes in faecal
samples of farm dogs and a cat. Parasite (Paris, France) 6, 189–193 (1999).
76. Dengjel, B. et al. Zoonotic potential of Enterocytozoon bieneusi. J Clin Microbiol 39, 4495–4499 (2001).
77. Santin, M., Cortes Vecino, J.A., and Fayer, R. Enterocytozoon bieneusi genotypes in dogs in Bogota,
Colombia. Am J Trop Med Hyg 79, 215–217 (2008).
78. Lores, B., del Aguila, C., and Arias, C. Enterocytozoon bieneusi (microsporidia) in faecal samples from
domestic animals from Galicia, Spain. Mem Inst Oswaldo Cruz 97, 941–945 (2002).
79. del Aguila, C. et al. Enterocytozoon bieneusi in animals: Rabbits and dogs as new hosts. J Eukaryot
Microbiol 46, S8–S9 (1999).
80. Breitenmoser, A.C., Mathis, A., Burgi, E., Weber, R., and Deplazes, P. High prevalence of Enterocytozoon
bieneusi in swine with four genotypes that differ from those identified in humans. Parasitology 118(Pt 5),
447–453 (1999).
81. Santin, M., Vecino, J.A., and Fayer, R. A zoonotic genotype of Enterocytozoon bieneusi in horses.
J Parasitol 96, 157–161 (2010).
Biology of Foodborne Parasites
82. Wagnerova, P. et al. Enterocytozoon bieneusi and Encephalitozoon cuniculi in horses kept under different
management systems in the Czech Republic. Vet Parasitol 190, 573–577 (2012).
83. Rinder, H. et al. Close genotypic relationship between Enterocytozoon bieneusi from humans and pigs
and first detection in cattle. J Parasitol 86, 185–188 (2000).
84. Santin, M. and Fayer, R. A longitudinal study of Enterocytozoon bieneusi in dairy cattle. Parasitol Res
105, 141–144 (2009).
85. Santin, M., Dargatz, D., and Fayer, R. Prevalence and genotypes of Enterocytozoon bieneusi in weaned
beef calves on cow-calf operations in the USA. Parasitol Res 110, 2033–2041 (2012).
86. Del Coco, V.F. et al. First report of Enterocytozoon bieneusi from dairy cattle in Argentina. Vet Parasitol
199, 112–115 (2014).
87. Fayer, R., Santin, M., and Trout, J.M. Enterocytozoon bieneusi in mature dairy cattle on farms in the
eastern United States. Parasitol Res 102, 15–20 (2007).
88. Santin, M., Trout, J.M., and Fayer, R. Enterocytozoon bieneusi genotypes in dairy cattle in the eastern
United States. Parasitol Res 97, 535–538 (2005).
89. Santin, M., Trout, J.M., and Fayer, R. Prevalence of Enterocytozoon bieneusi in post-weaned dairy calves
in the eastern United States. Parasitol Res 93, 287–289 (2004).
90. Fayer, R., Santin, M., and Trout, J.M. First detection of microsporidia in dairy calves in North America.
Parasitol Res 90, 383–386 (2003).
91. Lee, J.H. Molecular detection of Enterocytozoon bieneusi and identification of a potentially humanpathogenic genotype in milk. Appl Environ Microbiol 74, 1664–1666 (2008).
92. Deplazes, P., Mathis, A., Muller, C., and Weber, R. Molecular epidemiology of Encephalitozoon cuniculi
and first detection of Enterocytozoon bieneusi in faecal samples of pigs. J Eukaryot Microbiol 43, 93S
93. Buckholt, M.A., Lee, J.H., and Tzipori, S. Prevalence of Enterocytozoon bieneusi in swine: An 18-month
survey at a slaughterhouse in Massachusetts. Appl Environ Microbiol 68, 2595–2599 (2002).
94. Sak, B., Kvac, M., Hanzlikova, D., and Cama, V. First report of Enterocytozoon bieneusi infection on a
pig farm in the Czech Republic. Vet Parasitol 153, 220–224 (2008).
95. Li, W. et al. High diversity of human-pathogenic Enterocytozoon bieneusi genotypes in swine in northeast China. Parasitol Res 113, 1147–1153 (2014).
96. Jeong, D.K. et al. Occurrence and genotypic characteristics of Enterocytozoon bieneusi in pigs with diarrhea. Parasitol Res 102, 123–128 (2007).
97. Reetz, J. et al. First detection of the microsporidium Enterocytozoon bieneusi in non-mammalian hosts
(chickens). Int J Parasitol 32, 785–787 (2002).
98. Kasickova, D., Sak, B., Kvac, M., and Ditrich, O. Sources of potentially infectious human microsporidia:
Molecular characterisation of microsporidia isolates from exotic birds in the Czech Republic, prevalence
study and importance of birds in epidemiology of the human microsporidial infections. Vet Parasitol 165,
125–130 (2009).
99. Lobo, M.L. et al. Identification of potentially human-pathogenic Enterocytozoon bieneusi genotypes in
various birds. Appl Environ Microbiol 72, 7380–7382 (2006).
100. Lallo, M.A., Calabria, P., and Milanelo, L. Encephalitozoon and Enterocytozoon (Microsporidia) spores
in stool from pigeons and exotic birds: Microsporidia spores in birds. Vet Parasitol 190, 418–422 (2012).
101. Haro, M., Izquierdo, F., Henriques-Gil, N., Andres, I., Alonso, F., Fenoy, S., and del Aguila, C. First
detection and genotyping of human-associated Microsporidia in pigeons from urban parks. Appl Environ
Microbiol 71, 3153–3157 (2005).
102. Bart, A., Wentink-Bonnema, E.M., Heddema, E.R., Buijs, J., and van Gool, T. Frequent occurrence of
human-associated microsporidia in fecal droppings of urban pigeons in Amsterdam, the Netherlands.
Appl Environ Microbiol 74, 7056–7058 (2008).
103. Graczyk, T.K. et al. Urban feral pigeons (Columba livia) as a source for air- and waterborne contamination with Enterocytozoon bieneusi spores. Appl Environ Microbiol 73, 4357–4358 (2007).
104. Pirestani, M., Sadraei, J., and Forouzandeh, M. Molecular characterization and genotyping of human
related microsporidia in free-ranging and captive pigeons of Tehran, Iran. Infect Genet Evol 20, 495–499
105. Muller, M.G., Kinne, J., Schuster, R.K., and Walochnik, J. Outbreak of microsporidiosis caused by
Enterocytozoon bieneusi in falcons. Vet Parasitol 152, 67–78 (2008).
Enterocytozoon bieneusi
106. Guo, Y. et al. Host specificity and source of Enterocytozoon bieneusi genotypes in a drinking source
watershed. Appl Environ Microbiol 80, 218–225 (2014).
107. Nemejc, K. et al. Prevalence and diversity of Encephalitozoon spp. and Enterocytozoon bieneusi in wild
boars (Sus scrofa) in Central Europe. Parasitol Res 113, 761–767 (2014).
108. Dowd, S., Gerba, C., and Pepper, I. Confirmation of the human pathogenic Microsporidia Enterocytozoon
bieneusi, Encephalitozoon intestinalis, and Vittaforma corneae in water. Appl Environ Microbiol 64,
3332–3335 (1998).
109. Thurston-Enriquez, J. et al. Detection of protozoan parasites and microsporidia in irrigation waters used
for crop production. J Food Prot 65, 378–382 (2002).
110. Coupe, S. et al. Detection of Cryptosporidium, Giardia and Enterocytozoon bieneusi in surface water,
including recreational areas: A one-year prospective study. FEMS Immunol Med Microbiol 47, 351–359
111. Graczyk, T.K. et al. Human zoonotic enteropathogens in a constructed free-surface flow wetland.
Parasitol Res 105, 423–428 (2009).
112. Galvan, A.L. et al. Molecular characterization of human-pathogenic microsporidia and Cyclospora cayetanensis isolated from various water sources in Spain: A year-long longitudinal study. Appl Environ
Microbiol 79, 449–459 (2013).
113. Ben Ayed, L. et al. Survey and genetic characterization of wastewater in Tunisia for Cryptosporidium
spp., Giardia duodenalis, Enterocytozoon bieneusi, Cyclospora cayetanensis and Eimeria spp. J Water
Health 10, 431–444 (2012).
114. Wolk, D.M. et al. A spore counting method and cell culture model for chlorine disinfection studies of
Encephalitozoon syn. Septata intestinalis. Appl Environ Microbiol 66, 1266–1273 (2000).
115. Fournier, S. et al. Detection of microsporidia, cryptosporidia and giardia in swimming pools: A one year
prospective study (see Crypto library). FEMS Immunol Med Microbiol 33, 209–213 (2002).
116. Sodqi, M. et al. Unusual pulmonary Enterocytozoon bieneusi microsporidiosis in an AIDS patient: Case
report and review. Scand J Infect Dis 36, 230–231 (2004).
117. Botterel, F., Minozzi, C., Vittecoq, D., and Bouree, P. Pulmonary localization of Enterocytozoon bieneusi
in an AIDS patient: Case report and review. J Clin Microbiol 40, 4800–4801 (2002).
118. Weber, R. et al. Pulmonary and intestinal microsporidiosis in a patient with the acquired immunodeficiency syndrome. Am Rev Respir Dis 146, 1603–1605 (1992).
119. Brasil, P. et al. Clinical and diagnostic aspects of intestinal microsporidiosis in HIV-infected patients with
chronic diarrhea in Rio de Janeiro, Brazil. Rev Inst Med Trop Sao Paulo 42, 299–304 (2000).
120. Mor, S.M., Tumwine, J.K., Naumova, E.N., Ndeezi, G., and Tzipori, S. Microsporidiosis and malnutrition in children with persistent diarrhea, Uganda. Emerg Infect Dis 15, 49–52 (2009).
121. Velásquez, J.N. et al. Molecular identification of protozoa causing AIDS-associated cholangiopathy in
Buenos Aires, Argentina. Acta Gastroenterol Latinoam 42, 301–308 (2012).
122. Pol, S. et al. Microsporidia infection in patients with the human immunodeficiency virus and unexplained
cholangitis. N Engl J Med 328, 95–99 (1993).
123. Carr, A., Marriott, D., Field, A., Vasak, E., and Cooper, D.A. Treatment of HIV-1-associated microsporidiosis and cryptosporidiosis with combination antiretroviral therapy. Lancet 351, 256–261 (1998).
124. Guerard, A. et al. Intestinal microsporidiosis occurring in two renal transplant recipients treated with
mycophenolate mofetil. Transplantation 68, 699–707 (1999).
125. Conteas, C.N., Berlin, O.G., Ash, L.R., and Pruthi, J.S. Therapy for human gastrointestinal microsporidiosis. Am J Trop Med Hyg 63, 121–127 (2000).
126. Goetz, M., Eichenlaub, S., Pape, G.R., and Hoffmann, R.M. Chronic diarrhea as a result of intestinal
microsposidiosis in a liver transplant recipient. Transplantation 71, 334–337 (2001).
127. Molina, J.M. et al. Fumagillin treatment of intestinal microsporidiosis. N Engl J Med 346, 1963–1969
128. Audemard, A. et al. Fumagillin-induced aseptic meningoencephalitis in a kidney transplant recipient
with microsporidiosis. Transpl Infect Dis 14, E147–E149 (2012).
129. Sadler, F. et al. Genotyping of Enterocytozoon bieneusi in AIDS patients from the north west of England.
J Infect 44, 39–42 (2002).
130. ten Hove, R.J. et al. Characterization of genotypes of Enterocytozoon bieneusi in immunosuppressed and
immunocompetent patient groups. J Eukaryot Microbiol 56, 388–393 (2009).
Biology of Foodborne Parasites
131. Liguory, O., Sarfati, C., Derouin, F., and Molina, J.M. Evidence of different Enterocytozoon bieneusi
genotypes in patients with and without human immunodeficiency virus infection. J Clin Microbiol 39,
2672–2674 (2001).
132. Liguory, O., David, F., Sarfati, C., Derouin, F., and Molina, J.M. Determination of types of Enterocytozoon
bieneusi strains isolated from patients with intestinal microsporidiosis. J Clin Microbiol 36, 1882–1885
133. Rinder, H., Katzwinkel-Wladarsch, S., and Loscher, T. Evidence for the existence of genetically distinct
strains of Enterocytozoon bieneusi. Parasitol Res 83, 670–672 (1997).
134. Stark, D. et al. Limited genetic diversity among genotypes of Enterocytozoon bieneusi strains isolated
from HIV-infected patients from Sydney, Australia. J Med Microbiol 58, 355–357 (2009).
135. Chabchoub, N. et al. Genotype identification of Enterocytozoon bieneusi isolates from stool samples of
HIV-infected Tunisian patients. Parasite (Paris, France) 19, 147–151 (2012).
136. Ojuromi, O.T. et al. Identification and characterization of microsporidia from fecal samples of HIVpositive patients from Lagos, Nigeria. PLoS ONE 7, e35239 (2012).
137. Leelayoova, S. et al. Identification of genotypes of Enterocytozoon bieneusi from stool samples from
human immunodeficiency virus-infected patients in Thailand. J Clin Microbiol 44, 3001–3004 (2006).
138. Wumba, R. et al. Intestinal parasites infections in hospitalized AIDS patients in Kinshasa, Democratic
Republic of Congo. Parasite (Paris, France) 17, 321–328 (2010).
139. Wang, L. et al. Concurrent infections of Giardia duodenalis, Enterocytozoon bieneusi, and Clostridium
difficile in children during a cryptosporidiosis outbreak in a pediatric hospital in China. PLoS Neglect
Trop Dis 7, e2437 (2013).
140. Halanova, M. et al. Occurrence of microsporidia as emerging pathogens in Slovak Roma children and
their impact on public health. Ann Agric Environ Med 20, 695–698 (2013).
141. Ye, J. et al. Occurrence of human-pathogenic Enterocytozoon bieneusi, Giardia duodenalis and
Cryptosporidium genotypes in laboratory macaques in Guangxi, China. Parasitol Int 63, 132–137 (2014).
142. Abe, N. and Kimata, I. Molecular survey of Enterocytozoon bieneusi in a Japanese porcine population.
Vector Borne Zoonotic Dis (Larchmont, NY) 10, 425–427 (2010).
143. Reetz, J. et al. Identification of Encephalitozoon cuniculi genotype III and two novel genotypes of
Enterocytozoon bieneusi in swine. Parasitol Int 58, 285–292 (2009).
144. Ayinmode, A.B., Ojuromi, O.T., and Xiao, L. Molecular identification of Enterocytozoon bieneusi isolates from Nigerian children. J Parasitol Res 2011, 129542 (2011).
145. Sulaiman, I.M., Fayer, R., Lal, A.A., Trout, J.M., Schaefer III, F.W., and Xiao, L. Molecular characterization of Microsporidia indicates that wild mammals harbor host-adapted Enterocytozoon spp. as well as
human-pathogenic Enterocytozoon bieneusi. Appl Environ Microbiol 69, 4495–4501 (2003).
146. Zhao, W. et al. High prevalence of Enterocytozoon bieneusi in asymptomatic pigs and assessment of
zoonotic risk at a genotype level. Appl Environ Microbiol 80, 3699–3707 (2014).
147. Saksirisampant, W. et al. Intestinal parasitic infections: Prevalences in HIV/AIDS patients in a Thai
AIDS-care centre. Ann Trop Med Parasitol 103, 573–581 (2009).
148. Sokolova, O.I. et al. Emerging microsporidian infections in Russian HIV-infected patients. J Clin
Microbiol 49, 2102–2108 (2011).
149. Kicia, M. et al. Concurrent infection of the urinary tract with Encephalitozoon cuniculi and Enterocytozoon
bieneusi in a renal transplant recipient. J Clin Microbiol 52, 1780–1782 (2014).
150. Lee, J.H. Prevalence and molecular characteristics of Enterocytozoon bieneusi in cattle in Korea.
Parasitol Res 101, 391–396 (2007).
151. Abu Samra, N., Thompson, P.N., Jori, F., Zhang, H., and Xiao, L. Enterocytozoon bieneusi at the wildlife/
livestock interface of the Kruger National Park, South Africa. Vet Parasitol 190, 587–590 (2012).
152. Sak, B., Kvac, M., Kvetonova, D., Albrecht, T., and Pialek, J. The first report on natural Enterocytozoon
bieneusi and Encephalitozoon spp. infections in wild East-European House Mice (Mus musculus musculus) and West-European House Mice (M. m. domesticus) in a hybrid zone across the Czech RepublicGermany border. Vet Parasitol 178, 246–250 (2011).
153. Fayer, R., Santin, M., and Macarisin, D. Detection of concurrent infection of dairy cattle with Blastocystis,
Cryptosporidium, Giardia, and Enterocytozoon by molecular and microscopic methods. Parasitol Res
111, 1349–1355 (2012).
154. Jurankova, J., Kamler, M., Kovarcik, K., and Koudela, B. Enterocytozoon bieneusi in Bovine Viral
Diarrhea Virus (BVDV) infected and noninfected cattle herds. Res Vet Sci 94, 100–104 (2013).
Simone M. Cacciò and Marco Lalle
10.1 Introduction....................................................................................................................................175
10.2 Morphology and Classification......................................................................................................175
10.3 Biology, Genetics, and Genomics..................................................................................................178
10.4 Diagnosis and Typing.................................................................................................................... 180
10.5 Epidemiology and Molecular Epidemiology.................................................................................182
10.6 Pathogenesis and Clinical Features...............................................................................................186
10.7 Treatment and Prevention..............................................................................................................188
References............................................................................................................................................... 190
10.1 Introduction
The protozoan flagellate Giardia duodenalis is the etiological agent of giardiasis, one of the commonest
gastrointestinal infections of mammals, including humans, with a worldwide distribution. The infection
is transmitted by the fecal–oral route through ingestion of cysts, by both direct and indirect routes. In
humans, infection is caused by two genetically distinct groups of G. duodenalis, namely, assemblages A
and B, whereas the remaining six assemblages (C through H) described to date infect other mammals,
with various degrees of host specificity. Humans acquire infections mostly by consumption of contaminated water, and the numerous outbreaks reported further underline the important role that water plays
in the transmission of Giardia. In comparison, less is known about foodborne giardiasis, due to the
difficulties in investigation of cases and to the lack of standard methods for the detection of Giardia on
foodstuffs. Clinical giardiasis occurs only in a percentage of infected individuals, whereas many cases
are asymptomatic, but the reasons for this variability in clinical presentation are unknown. In recent
years, important progress has been made in understanding the taxonomy, genetics, epidemiology, and
pathogenesis of this organism. This chapter highlights relevant advances in these fields.
10.2 Morphology and Classification
The life cycle of Giardia is simple and comprises only two stages: the trophozoite, a noninvasive form
that replicates actively on the mucosa of the small intestine, and the environmentally resistant cyst,
which represents the transmittable stage1 (Figure 10.1).
The G. duodenalis trophozoite (Figure 10.2) is a pear-shaped cell about 12–15 μm long, 6–8 μm
wide, and 1–2 μm thick (see Table 10.1 for morphological features of the other species). Trophozoites
contain two nuclei surrounded by nuclear envelopes and have a complex cytoskeleton that maintains
their shape and anchors the four pairs of flagella, the median body, and the ventral disk.1 The flagella
are composed of microtubules in a typically eukaryotic 9 + 2 arrangement and are built from basal
bodies located between the nuclei. The median bodies are formed by an irregular set of microtubules,
Biology of Foodborne Parasites
FIGURE 10.1 Schematic representation of the life cycle of Giardia.
Anterior flagella
Cyst wall
Adhesive disc
Basal body
Median bodies
Disc fragments
Peripheral vesicles
Ventral flagella
Caudal flagella
Peripheral vesicles
Flagellar axonemes
FIGURE 10.2 Graphical representation of the structure of (a) a trophozoite and (b) a cyst.
have a comma shape structure that varies in size and thickness, and are located transversally, perpendicular to the central axis. To date, no exact function has been assigned to the median bodies, but a
role as a reserve of microtubules or in the biogenesis of the ventral disk has been proposed. The ventral
disk is a unique structure of Giardia and covers the anterior half of the ventral side of the trophozoite.
It is linked to the plasma membrane by short fibers that are composed of α- and β-tubulins, contractile
proteins, and cytoskeletal proteins called giardins. The ventral disk is essential for the attachment of
the trophozoite to the enterocytes and is thought to play roles also in nuclear division. This structure
undergoes profound changes during the process of differentiation into cysts where it disassembles and
is stored as fragments during cyst formation.2 Giardia has a well-developed endoplasmic reticulum
and can form secretory vesicles, some of which are clearly visible during encystation (the encystationspecific vesicles, ESVs) and transport of cyst wall components to the cell surface for assembly into the
extracellular cyst wall.3
Surprising, a typical Golgi apparatus comprising a set of flattened cisternae is not found at any time
during the life cycle. Giardia also lacks other “typical” eukaryotic organelles, like peroxisomes and
mitochondria, although a mitochondrial remnant has been described and recently shown to be involved
in FeS cluster assembly.4 In the cell cytoplasm and adjacent to the plasma membrane (except in the region
of the ventral disk), there are numerous vesicles and tubules, the peripheral vacuoles, that function both
as endosomes and lysosomes.5
TABLE 10.1
Six Species Currently Recognized within the Giardia Genus, Their Host Distribution,
and Distinctive Morphological Characteristics
Distinctive Morphological Features
G. muris
Humans, domestic
and wild mammals
G. microti
G. ardeae
G. psittaci
G. agilis
Pear-shaped trophozoites with
claw-shaped median bodies.
Rounded trophozoites with small,
round median bodies.
Trophozoites similar to G. duodenalis.
Cysts contain fully differentiated
Rounded trophozoites, prominent
notch in ventral disk and rudimentary
caudal flagellum. Median bodies
round–oval to claw shaped.
Pear-shaped trophozoites, with no
ventrolateral flange. Claw-shaped
median bodies.
Long and narrow trophozoites with
club-shaped median bodies.
G. duodenalis
Length/Width of the
Trophozoite (μm)
The cysts of Giardia are oval in shape and range in size from 6 to 10 μm. The cyst wall is about
0.3–0.5 μm in thickness and is formed by an outer filamentous layer and an inner membranous layer
including two membranes that enclose the periplasmic space. The cyst wall is composed of carbohydrates,
in the form of N-acetylgalactosamine polymers, and cyst wall proteins. The cytoplasm of the mature cyst
contains four nuclei, the contracted flagella, and fragmented portions of the ventral disk (Figure 10.2).
On the basis of morphological characters, organisms in the genus Giardia are classified in the phylum
Metamonada, subphylum Trichozoa, superclass Eopharyngia, class Trepomonadea, subclass Diplozoa,
and order Giardiida.6 The taxonomy of Giardia species has been, and still is, a subject of intense debate
and controversy, and this has resulted in a confusing nomenclature with different names being used for
the same species.7 Early taxonomy was based on the assumption of strict host specificity (i.e., a different species for each host), and a total of 51 species of Giardia have been described, including 30 from
mammals (of which 2 were from humans), 14 from birds, 4 from amphibians, 2 from reptiles, and 1 from
fish.6 It was not until the seminal work of Filice, however, that the taxonomy of Giardia species was
reconsidered.6 Filice concluded that no experimental evidence supported the validity of species based
on host specificity and, therefore, rejected this criterion. He proposed to consider only three morphologically distinct groups, namely, Giardia muris, Giardia agilis, and G. duodenalis, mainly on the basis
of the shape of internal median bodies as well as body shape and length. This scheme rapidly became
accepted by the scientific community, albeit, as acknowledged by Filice himself, many species were put
under the G. duodenalis “umbrella,” due to the lack of tools to discriminate reliably between variants.
With the development of advanced microscopic techniques, ultrastructural description of trophozoites
allowed the description of two new species from birds, Giardia ardeae and Giardia psittaci, whereas
Giardia microti, a parasite infecting various rodents, was recognized as a separate species due to the
unique presence of fully differentiated trophozoites in the cysts. Therefore, six species are recognized in
the genus (Table 10.1).
The development of techniques for the in vitro propagation of trophozoites in axenic conditions8
opened the possibility to characterize Giardia strains from different hosts using genetic techniques.
The first important series of experiments aimed at comparing strains on the basis of polymorphisms
of many isoenzymes and revealed a large amount of genetic variability among G. duodenalis strains.6
Importantly, clustering analysis of the isoenzyme data identified strongly supported groups of genetically related strains that, in most cases, were derived from specific hosts.9 Those genetic groups are
nowadays referred to as assemblages.
Biology of Foodborne Parasites
TABLE 10.2
Currently Recognized G. duodenalis Assemblages, Their Host Distribution,
and Their Previously Proposed Taxonomy
Host Distribution
Proposed Species Name
Humans and other primates, livestock, dogs,
cats, and some species of wild mammals
Humans and other primates, dogs, cats, and
some species of wild mammals
Dogs and other canids
Dogs and other canids
Hoofed livestock
Marine mammals (pinnipeds)
G. duodenalis
G. enterica
G. canis
G. bovis
G. cati
G. simondi
In more recent years, sequencing and phylogenetic analysis of gene fragments amplified by polymerase chain reaction (PCR) from field isolates (i.e., not expanded by in vitro growth) further corroborated the validity of assemblages and helped to estimate the level of intra-assemblage genetic variability
by assessing mutation at synonymous positions, which are not detectable when protein polymorphisms
are studied.10,11 On the basis of molecular studies performed on parasites isolated from many hosts, eight
assemblages have been described, among which assemblages A and B infect humans and other animals,
assemblages C and D are restricted to carnivores, assemblage E to hoofed animals, assemblage F to
cats, assemblage G to rats, and assemblage H to marine mammals (Table 10.2). The genetic differences
separating the eight assemblages are very large and are paralleled by phenotypic differences in terms
of adaptability to axenic conditions and metabolism and susceptibility to drugs and to infection with a
dsRNA virus.6 Recent comparisons at the whole genome level have reinforced the notion that assemblages A, B, and E represent distinct species.12,13
Therefore, it has been proposed that the different assemblages deserve the status of species and that
previously assigned names are available for most assemblages6 (Table 10.2). However, controversial
points still remain, mostly concerning the names to be given to assemblages A and B (the proposed
name are G. duodenalis and Giardia enterica, respectively), to assemblage H, and the fact that the two
distinct assemblages (C and D) that infect dogs will be grouped in a single species, Giardia canis, despite
the rather large genetic variability observed at the genes investigated. It can be anticipated that further
genomic analyses of assemblages, and of genotypes within each assemblage, will contribute significantly
to the establishment of a robust taxonomy. Formal redescriptions of these species incorporating genetic
data are needed before these species names within G. duodenalis sensu lato can be used.
10.3 Biology, Genetics, and Genomics
Giardia has long been considered as a primitive, early diverging eukaryote, mainly because of the
absence of a typical Golgi apparatus, of peroxisomes, and of mitochondria, and therefore an important
model for evolutionary studies. While this view has been rejected on the basis of recent findings from
more accurate phylogenetic studies and genomic data,6 Giardia remains an interesting model system to
study the mechanisms of cellular differentiation in early branching eukaryotes.14 As mentioned earlier,
the life cycle of Giardia is simple; however, two major developmental transitions must occur, excystation and encystation. The excystation process starts after the ingestion of inert cysts by the host and is
initially triggered by host stomach acids; then the cyst passes into the small intestine before rupturing.
Flagella first appear through an opening in one of the poles of the cyst, followed by the excyzoite body.
Cysteine proteases, released from the lysosome-like peripheral vesicles, are thought to have an important role in this process by degrading the cyst wall from the inside. The excyzoite undergoes cytokinesis
twice without intervening S phases, finally producing four trophozoites. During the division process,
the excyzoite increases its metabolism and gene expression, segregates organelles, upregulates proteins
associated with motility, and assembles the adhesive disk. Studies of gene expression during excystation
have revealed that a number of genes involved in protein degradation, the cytoskeleton, and mitosis are
upregulated, in agreement with the fast reassembly of the cytoskeleton and the fast cell division seen
during early excystation.14
The process of encystation can be divided into three stages.15 The first is the reception of the stimulus for
encystation, the transmission of this signal to the nuclei, and the expression of encystation-specific genes.
The second step comprises the synthesis of precursors and cyst wall molecules, the biogenesis of secretory
organelles absent in non-encysting trophozoites, and the intracellular trafficking of the cyst wall components. The last stage of encystation includes exocytosis of ESVs and assembly of the extracellular cyst wall.
This process is induced in response to host-specific factors such as high levels of bile, low levels of
cholesterol, and a basic pH. Giardia cannot synthesize cholesterol de novo, and trophozoites take up
this lipid from the milieu of the upper small intestine.1 When the parasite travels down the intestine, the
concentration of cholesterol becomes low and this triggers encystation, albeit the mechanism used by
Giardia to detect low level of cholesterol is unknown.
The genome of Giardia has the features expected of eukaryotic cells, including linear chromosomes
flanked by telomeres that are similar in sequence (TAGGG) to those of other eukaryotes.1 Like other
diplomonads, Giardia has two nuclei that have been shown to be equivalent in size and in the amount of
DNA that they contain and are both transcriptionally active.
Giardia has a simplified transcriptional apparatus, but gene expression seems largely regulated at the
transcriptional level and RNA processing. Studies of transcription in G. duodenalis have employed serial
analysis of gene expression (SAGE) and oligonucleotide microarrays. SAGE was applied to characterize
10 time points throughout the life cycle and uncovered a certain extent of differential gene expression
throughout the life cycle. SAGE also indicated abundant antisense transcription.16 Promoters in G. duodenalis are suggested to be bidirectional, in part explaining the production of antisense transcripts. In
addition, promiscuous transcription driven by “cryptic” promoters, containing simple A+T-rich stretches
also contribute to antisense transcription. However, transcript discovery using SAGE is limited by
underrepresentation of genes with low expression levels, and this method has largely been replaced by
high-throughput sequencing assays. Oligonucleotide microarrays were used to identify gene expression
changes in drug-resistant clones,17 encysting cells and trophozoites during host–parasite interactions.18,19
The first genome of Giardia was published in 2007, after years of investigation, and was obtained from
genomic DNA extracted from a population of trophozoites of the WB strain of assemblage A, genotype
A1.20 As the title of the paper stated, the genome is “minimalist,” not only because of its small size
(about 11.7 Mb), the scarcity of introns, and the short length of intergenic sequences (average 352 bp) but
also because Giardia has genes for rudimentary forms of many cellular processes, with fewer subunits
present in simplified cellular machineries, including those involved in DNA synthesis and transcription, RNA processing, cytoskeletal function, and cell division. Further, Giardia has a limited metabolic
repertoire with many bacterial-like enzymes that were introduced by horizontal gene transfer.20 Protein
kinases comprised the largest class of proteins and are likely involved in the complex signal transduction network needed for parasite differentiation. Notably, most of the genes are of unknown function and
remained unverified by experimental or informatics means.
With the recent advent of the so-called next-generation sequencing (NGS) techniques, it is possible
to generate huge amounts of sequences at a much lower cost and in a shorter time, and this is having an
impact also on Giardia genomics. Indeed, two genome drafts were recently obtained using NGS12,13 and
have provided data for a human-derived isolate of assemblage B (the GS strain) and a pig-derived isolate
from the hoofed-specific assemblage E (the P15 strain). Comparative genomics has revealed that the
G. duodenalis genome consists of a “core” of conserved genes (approximately 90% of the genes) and a
more variable part of the genome (about 9%) that contains members of large gene families such as variable surface proteins (VSPs), NEK kinases, protein 21.1, and high cysteine membrane protein. A limited
number of genes (not part of the large gene families) appear to be present in just one assemblage (5 in
WB, 31 in GS, and 38 in P15). Most of these genes have no sequence similarity to known sequences, and
their functions are, therefore, unknown; some have been acquired recently from bacteria.13 Comparative
genomics have also detected many structural rearrangements confirming that the G. duodenalis genome
Biology of Foodborne Parasites
is endowed with considerable plasticity.13 Plasticity is mainly observed at subtelomeric regions of the
chromosomes, in agreement with studies based on pulsed-field gel electrophoresis that have shown considerable size variation,1 and this is due to differences in the number of rDNA molecules and the presence of retroposons, pseudogenes, and low-complexity repeats. The internal regions of the chromosomes
correspond to the more stable conserved “core” genome and show a much higher level of synteny among
genomes, but polymorphisms (both at synonymous and nonsynonymous positions) and deletions/insertions are commonly observed in this part of the genome as well.
A striking difference between assemblage A and B genomes is the high level of allelic sequence heterozygosity (ASH) observed in the assemblage B, GS strain, which is almost completely lacking in the
assemblage A, WB strain.12 The finding of a very low level of ASH in the WB genome was surprising,
since G. duodenalis is supposed to reproduce asexually and is thus expected to accumulate independent
mutations in the two nuclei.20 Trophozoites have a ploidy of four, with two sets of five chromosomes in
each nucleus, and replication is equational rather than reductional, which means that nuclear asymmetry
is maintained throughout the replication cycle.21 Therefore, if nuclei are asexual, they would expect to
diverge one from another, but the low ASH indicates that alleles within nuclei as well as those from
opposite nuclei are nearly identical, suggesting the presence of homogenization mechanisms. There is
now evidence that genetic exchange occurs between G. duodenalis isolates of assemblage A, genotype
AII22 and some data suggest that this also occurs between assemblage A and B isolates.23 Furthermore,
the Giardia genome contains many (21 of 29) of the genes required for meiosis,24 albeit their exact role is
unknown, as they can be involved in other functions, such as DNA repair. Thus, whether a true meiotic
process occurs in G. duodenalis remains to be established. Based on experimental evidence of plasmid
transfer between nuclei in the cyst, and fusion between cyst nuclei, it has been proposed that somatic
homologous recombination can take place during encystation and that the process is facilitated by the
meiosis gene homologs, whose expression was observed in cysts but not in trophozoites.25 Therefore, a
body of data suggest that genetic exchange occurs in G. duodenalis, even if the exact mechanisms remain
elusive.26 Knowledge about the mechanisms and frequency of genetic recombination will be very important also to understand the spread of genes involved in pathogenicity and drug susceptibility.
Therefore, there are several important biological questions that can be addressed by further genomic
studies. The current drafts of the genomes of assemblages B and E were assembled from sequences
obtained by pyrosequencing, a methodology that is poor at covering regions containing repetitive DNA,
which in G. duodenalis mainly corresponds to the subtelomeric portions of the chromosomes and to
genes that contain conserved repetitive domains (like VSP). Therefore, those drafts are fragmented into a
high number of contigs and, while suitable for insightful genomic comparisons, good reference genomes
are needed for all G. duodenalis assemblages, particularly for assemblage B.
10.4 Diagnosis and Typing
The diagnosis of Giardia is mainly based on the microscopic demonstration of cysts in fecal samples,
but immunological and molecular assays are also available and increasingly used.27 Fecal suspension
or concentrates are used to prepare wet mounts and then examined either unstained or stained (the primary stain for cysts is iodine). When possible, fixation in sodium acetate–acetic acid–formalin (SAF) is
recommended, as this increases the detection rate of intestinal protozoa, including Giardia, compared
to unfixed stools. Due to the intermittent excretion of cysts, at least three stool specimens should be
examined for a definitive diagnosis. The sensitivity of microscopy is rather low, and competent microscopists are needed to recognize Giardia cysts and to distinguish them from other organisms present in
the feces having a similar morphology. Identification relies on characteristic morphological features
of the cyst. These include shape (ellipsoid to oval, with smoothed wall, colorless, and refractile), size
(8–14 × 6–10 μm), and internal structures (presence of four nuclei at a pole of the cyst, flagellar axonemes
that lie diagonally across the long axis of the cyst, and fragments of the ventral disk as “crescent bodies”;
see Figure 10.2).
Several immunologic-based assays have been developed for the diagnosis of Giardia.28 A widely
used assay is immunofluorescence with monoclonal antibodies (mAbs) that recognize surface-exposed
epitopes of the cyst. The antibodies are usually conjugated with fluorescein isothiocyanate, which fluoresces green when examined with a fluorescence microscope using the blue filter block. Various commercial kits are available to perform immunofluorescence.
Another diagnostic method is based on the detection of antigens in feces, and the preferred format is
the antigen capture ELISA test, which utilizes antibodies to capture antigens on the solid phase and to
detect its presence by developing a colored reaction product. The target antigen could be as complex as
an aqueous extract of trophozoites or an individual molecule, such as the Giardia stool antigen (GSA),
an antigen of a molecular mass of 65 kDa, which is present in both trophozoites and cysts. The sensitivity and specificity of ELISA are comparable or better than that of microscopy. When performed using
GSA, the ELISA can reach a sensitivity of 95%–100% and a specificity of 100%.27 In addition, ELISA
simplifies the screening of a large number of samples, for example, during a suspected outbreak, and
the clinical follow-up of patients after treatment. The use of lateral flow immunochromatography assay
(­dipstick) considerably reduces the time needed for antigen–antibody interaction and can generate results
in 15 min, providing a convenient alternative method for detecting Giardia antigens in stool samples.
The reaction takes place in a cassette where soluble parasite antigens can bind to immobilized antibodies
and generates a qualitative result seen as a colored band at a specific location marked on the cassette.
Detection of circulating antibodies can also be used as a diagnostic tool. It has been shown that 81%
of symptomatic individuals have IgG antibodies directed toward surface antigens of trophozoites, compared to 12% of controls.29 The level of IgG remains detectable for up to 18 months after drug treatment,
whereas that of IgM rapidly falls to control levels after drug treatment and can only be used as an indicator of current infection. Both ELISA and immunofluorescence can be used to detect IgG and IgA in the
serum of infected individuals. The main problem with serological assays is the difficulty in distinguishing the responses obtained from symptomatic and asymptomatic individuals. A good candidate is alphagiardin, an immunodominant antigen, which is strongly recognized by the sera of infected individuals,
as demonstrated by a study after an outbreak in Sweden.30
Molecular diagnostics tools are also available and are thought to provide higher sensitivity and specificity compared to both microscopic and immunologic assays.27 The main targets used for PCR-based
detection and characterization of Giardia are discussed below (see also Table 10.3). Conventional PCR,
either as a single or nested procedure followed by sequencing of amplification products, is widely used
for research purposes (e.g., epidemiological surveys). However, difficulties in the extraction of inhibitorfree DNA, the risk of cross-contamination, and, more generally, the lack of standardized methods limit
the applicability of PCR in routine diagnostic laboratories. In this context, the use of real-time PCR
procedures can reduce the risk of cross-contamination and, since this methodology can be tailored for
the simultaneous detection of multiple pathogens, it may represent a feasible diagnostic alternative in the
clinical laboratory.31
As mentioned earlier, many insights into the taxonomy and genetics of Giardia were obtained by
the study of protein polymorphisms, but this technique requires large amounts of material that can be
obtained only by in vitro culturing of parasites. It is well known that isolates of Giardia differ in their
ability to adapt to in vitro growth conditions and that those from assemblages B, C, and D are particularly refractory to this process.1
The introduction of molecular techniques, in particular those based on the in vitro amplification of
nucleic acids (i.e., PCR and related methodologies), has revolutionized the study of the epidemiology
of giardiasis. The first PCR assays targeted fragments of well-conserved eukaryotic genes, sometimes
using degenerated primers (18S small subunit ribosomal, glutamate dehydrogenase, elongation factor
1-α, triosephosphate isomerase),10 or genes uniquely associated with the parasite (beta-giardin).32 In more
recent studies, a number of other PCR assays have been developed and tested for their applicability for
detection and typing of G. duodenalis; a list of the currently available markers is given in Table 10.3.
It is important to recall that these markers differ widely in terms of genetic variability33; indeed,
some, like the 18S rDNA and the elongation factor 1-α, are strongly conserved and can be used to identify G. duodenalis assemblages but are of little use for studies where genetic variation within assemblages needs to be determined. Nevertheless, due to the multicopy nature of the 18S rDNA, the PCR
that targets this locus has a high sensitivity and is often used for detection of Giardia from different
matrices (e.g., human and animal feces, water samples). Some researchers have considered that Giardia
Biology of Foodborne Parasites
TABLE 10.3
List of the Genetic Loci Used for Genotyping, Their Function, and Availability of Information
from Different Giardia Species or G. duodenalis Assemblages
Genetic Marker
Sequences Availability
Glutamate dehydrogenase
Triosephosphate isomerase
Elongation factor 1-α
Histone H2B
Histone H4
Chaperonin 60
Open reading frame C4
18S rDNA
Involved in DNA repair
Housekeeping enzyme
Housekeeping enzyme
Structural protein
Involved in translation
Mediates electron transfer
Nucleosomal protein
Nucleosomal protein
Structural protein
Structural protein
Heat shock protein
Hypothetical heat shock protein
Small subunit of the ribosome
Intergenic ribosomal spacer
ITS1, ITS2, and 5.8S rDNA
Ribosomal protein L7a
Noncoding ribosomal
G. duodenalis assemblages A and B
G. duodenalis assemblages, G. muris, G. ardeae
G. duodenalis assemblages, G. microti, G. muris, G. ardeae
G. duodenalis assemblages, G. muris
G. duodenalis assemblages, G. muris, G. ardeae
G. duodenalis assemblages A and B
G. duodenalis assemblages A and B
G. duodenalis assemblages A and B
G. duodenalis assemblages A and B, G. ardeae
G. duodenalis assemblages A and B
G. duodenalis assemblages A and B
G. duodenalis assemblages A and B
G. duodenalis assemblages, G. muris, G. microti,
G. ardeae, G. agilis
G. duodenalis assemblages A and B
G. duodenalis assemblages, G. muris, G. microti, G. ardeae
G. duodenalis assemblages A and B
has a clonal population structure, and that the use of a single marker with high genetic variability can
provide a resolution as high as multilocus sequence typing (MLST).34 In more recent studies, however,
the use of a multilocus typing scheme has been shown to represent a more informative approach for
genotyping this parasite.35,36 Current MLST schemes are predominantly based on housekeeping genes,
and the variation found in these markers seems sufficient for genotyping. Highly variable molecular
markers for subtyping, such as microsatellites, which are necessary for tracing outbreaks, local transmission routes, and population genetics studies, have not been identified thus far and appear rarely
represented in the genome.
10.5 Epidemiology and Molecular Epidemiology
The epidemiology of giardiasis is complex. Infection occurs by any route by which material contaminated with cysts excreted by infected hosts can reach the mouth of a susceptible host. Therefore, both
direct and indirect transmission routes exist and the ability of cysts to persist in the environment accounts
for the important role of water and food in the epidemiology of giardiasis.37
The parasite has a global distribution, but the prevalence of infection is clearly higher in developing
regions of the world, where Giardia is common in both children and adults. In recognition of the burden of disease caused by Giardia and to underline its link to poverty, the WHO has included it in the
list of neglected disease since 2004.38 The majority of studies have focused on asymptomatic children,
with reported infection rates between 8% and 30% in developing countries and between 1% and 8%
in industrialized countries.39 Those rates are probably higher in individuals with diarrhea. Despite the
availability of different diagnostic methods (see “Diagnosis”), the current epidemiological scenario is
largely influenced by the fact that many countries did not report any data, by the lack of monitoring programs, and by the high rate of asymptomatic carriage of Giardia in humans. All these data suggest that
giardiasis is strongly underdiagnosed and underreported.
In humans, giardiasis is mainly a pediatric infection, with the highest prevalence observed in children
aged 1–4 years. This pattern is found both in industrialized and developing countries and is thought
to be due to lower hygiene and higher susceptibility of children at the first exposure to the parasite.
A secondary peak is observed in adults aged 30–40 years; in this case, women represent the risk category,
likely because of direct transmission of Giardia from children to their mothers. Other risk groups include
institutionalized children, gay men, returning travelers, immigrant/refugees, and adopted children.37,40
Giardiasis is not considered a major cause of enteritis in HIV-infected patients, and it is not listed
among the opportunistic parasitic infections because no prolonged symptoms are observed and therapy
is independent of the patient’s immune status. The observed prevalence varies between 1.5% and 17.7%
in the few reports published.41 The symptoms of giardiasis in HIV-infected individuals appear to be
similar to, and no more severe than, those of giardiasis in HIV-negative individuals, with asymptomatic
infection occurring commonly in the presence of HIV.29 However, when CD4+ counts are reduced and
cause progressive immunosuppression, the risk of symptomatic Giardia infections increases, with a
tendency toward chronic diarrhea.
In the 2009–2010 report of giardiasis from the United States, a marked seasonality was observed,
confirming the trend seen in previous years, with a twofold increase of cases during the summer, coinciding with increased outdoor activities (e.g., swimming and camping).42 Similarly, a study in New Zealand
between 1996 and 2000 showed a significant seasonal variation in notification of giardiasis, with peaks
in late summer and early autumn.43 Data from European countries showed an average monthly incidence
of ~950 cases with an autumn peak in September to November of ~1350 cases.37 Bearing in mind the
delay between infection, development of symptoms, and submission of specimens, which may amount
to 5 weeks or more, this late summer/autumn peak probably represents an increase in infection during
the mid- to late summer months. Infections associated with travel and outdoor recreation may at least
partially explain this pattern.
Risk factors for giardiasis have been mainly studied in industrialized countries. Much of our knowledge derives from outbreak investigations, whereas few studies have focused on endemic or sporadic
giardiasis. Outbreaks are most frequently waterborne and are caused by drinking or recreational water
contamination, although other transmission routes have also been implicated.37 The routes of transmission for sporadic cases are poorly known, but recent case-control studies have identified the importance
of person-to-person spread, travel, contact with livestock, and potable and recreational water as risk
factors for sporadic disease.
A retrospective case-control study in rural New England looked at 171 patients and 684 age- and sexmatched control and identified the household use of shallow water sources as the main risk factor, followed by foreign travel, day care center exposure, and household case contact.44 A matched case-control
study in the United Kingdom (232 cases and 574 controls) identified swallowing water while swimming,
recreational freshwater contact, drinking treated tap water, and eating lettuce as positively and independently associated with infection.45 A case-control study in Germany included 120 laboratory-­confirmed
autochthonous Giardia cases with clinical manifestations (diarrhea, cramps, bloating) and 240 randomly selected controls from the local population registry matched by county of residence and age
group.46 Cases were more likely to be male, immunocompromised, and daily consumers of green salad.
Remarkably, contact with animals (pets/farm animals) and exposure to surface water (swimming/water
sports) were not associated with symptomatic disease. A case-control study in Italy, performed during
the Catholic Jubilee in 2000, to look for the effect of mass gathering on the transmission of giardiasis,
enrolled 52 cases and 72 controls, all residents of Rome.47 Multivariate analysis showed that traveling
abroad, exposure to surface water, and high educational levels were the main risk factors associated
with giardiasis. Having a maid from a high-prevalence country was independently associated, although
without statistical significance.
A study in Auckland, New Zealand, explored the risk of nappy changing by comparing 183 patients
with Giardia-positive stools with 336 age-matched controls identified randomly from the telephone
book.48 The risk of infection was significantly higher for housewives and nursing mothers compared with
other occupational groups. Physical contact with children wearing nappies showed a significant association with giardiasis. Nappy changing was associated with a fourfold increased risk, and giardiasis was
also associated strongly with childcare center attendance. Of these two factors, childcare center attendance and nappy changing, only nappy changing remained a significant risk for infection after logistic
regression.48 The studies mentioned are difficult to compare because of their different design and have
highlighted different risk factors, albeit the importance of direct contact, particularly with children, and
of indirect contact, particularly with water, are common findings.
Biology of Foodborne Parasites
The ability of Giardia to persist in the aquatic environment is of paramount importance for the transmission of infection. Cysts can survive for months in cold water while retaining, at least in part, their
infectivity. Furthermore, cysts are relatively resistant to chlorination at the levels usually employed in
water treatment, particularly at low temperatures and high pH. Finally, Giardia infects humans and a
wide range of animals, and all these hosts contribute to a massive contamination of the environment. It
is, therefore, not surprising that the occurrence of Giardia cysts in different water types has been repeatedly documented, mostly in North America and United Kingdom but also from other parts of the world,
mainly Asia.49 Concentration varies greatly depending on the type of water, being higher in surface water
(0.01–100 cysts/L) than in groundwater or in recreational water.49 Results are obviously influenced by the
efficiency of the method used in detecting Giardia cysts in that particular water type. In the last 20 years,
the majority of studies have used methods based on US-EPA Method 1623.49,50 In a recent review, it has
been reported that, of the 199 outbreaks caused by protozoa during the period of 2004–2010, 70 (35%)
were caused by Giardia.50 It is interesting to note that those outbreaks were all reported from developed
countries where detection and monitoring systems are more likely to be in place. The largest waterborne
outbreak has occurred in Bergen (Norway) in 2004, with a total of 1300 laboratory-confirmed cases, and
was likely caused by sewage leakage from a residential area with drainage toward the water source.51
The water treatment plant involved in this outbreak was one of the oldest in Norway, and the treatment in
place during the outbreak was just chlorination. Interestingly, investigation of this outbreak has revealed
that at least 5% of infected individuals developed persistent postinfectious fatigue syndrome that resulted
in high rates of occupational disability.52 Moreover, about 10% of the outbreak patients had persistent
symptoms, with mean disease duration of 7 months, and one-third of them suffered of chronic giardiasis
showing treatment failure even after 1–3 courses of treatment with metronidazole (MTZ).
The role of food in the epidemiology of giardiasis is not well understood. This is at least partially
explained by the fact that detection of Giardia cysts on foodstuffs is a technically challenging process,
due to the low number of parasites that may be present and to the wide differences in food matrices, which
requires the development of ad hoc methods.53 The methods used are based on four basic steps: elution of
cysts from the foodstuffs, concentration of the extract by centrifugation and separation of the cysts from
food materials by immunomagnetic separation, immunofluorescence staining of the cysts to allow their
visualization, and identification of cysts by microscopy. Recently, efforts have been made toward the
implementation and validation of standard methods for detection of Giardia cysts on/in foods.54
Foods that have been found naturally contaminated with Giardia cysts include fruits (strawberries),
vegetables (dill, lettuce, mung bean sprouts, and radish sprouts), and shellfish (species of oysters and
clams).53 This contamination is an important public health consideration because these products are frequently consumed raw without thermal processing. Voluntary consumption of mud or sand (geophagia),
which is particularly common in children and individuals with mental problems, may also contribute to
the transmission of Giardia.
There are fewer foodborne outbreaks documented than waterborne outbreaks, probably because of
the lack of appropriate tools and/or because cases are more widespread, appearing sporadic, rather than
being locked into water distribution networks, which are associated with outbreaks of disease.
Most foodborne outbreaks of giardiasis were ascribed to direct contamination by a food handler, but in
two instances, a role for zoonotic transmission was suggested, namely, the consumption of a Christmas
pudding contaminated with rodent feces and tripe soup made from the offal of an infected sheep.53 Only
one outbreak has been investigated using molecular tools. The outbreak involved 30 school and church
staff members who had lunch in the same restaurant in San Francisco, CA, in September 2001. Stool
testing of restaurant employees identified three asymptomatically infected food handlers, including a
cook, a waiter, and a counter assistant. Two formalin-fixed stool specimens from school employees were
identified as having G. duodenalis assemblage B.39
Molecular typing techniques have been extensively used to study the complex epidemiology of giardiasis, with a particular focus on some controversial aspects including zoonotic transmission, the occurrence of mixed infection in humans, the potential for genetic exchange between parasite isolates, and the
correlation between clinical symptoms and Giardia assemblages or genotypes6,37,39 (for this latter aspect,
see Pathogenesis and Clinical Features). However, results obtained thus far have not provided clear
answers to those questions, likely because of the complexity of the issues, differences in study design
(which make comparisons across studies difficult), and the lack of standardized methods for genotyping
this parasite. For the sake of clarity, data are presented and discussed separately for assemblages A and B.
Earlier studies based on protein polymorphisms demonstrated genetic variability between and among
human and animal isolates of assemblage A. In the most recent published study,9 four subgroups (AI, AII,
AIII, and AIV) were described by the analysis of 10 isolates at 23 genetic loci, and the host distribution
indicated that human isolates belong to subgroups AI and AII, while animal isolates belong to subgroups
AI, AIII, and AIV. Therefore, only subgroup AI seems to have zoonotic potential, whereas subgroup AII
seems to be human specific.
The correlation between subgroups defined by protein analysis and those identified by DNA analysis
was demonstrated10 on two reference strains, Ad-1 for subgroup AI and Ad-2 for subgroup AII, which
were analyzed at four genetic loci. Surprisingly, as no DNA information has been generated from the
isolates that defined subgroups AIII (cats) and AIV (cat, alpaca, guinea pig), the existence of those subgroups is still based on protein polymorphisms. A number of studies have focused on the comparative
analysis of genetic polymorphisms of housekeeping genes (in most cases using a single locus) between
human and animal isolates, with the aim of defining transmission routes and zoonotic potential of various
animals, mainly livestock and pets.11 This has led to the description of new variants (sometimes referred
to as genotypes or subgenotypes), some of which were found in different hosts, including humans. This
finding was interpreted as an indirect evidence for zoonotic potential. The amount of DNA sequences
has accumulated rapidly over time, allowing extensive evaluation of the genetic variability between and
within G. duodenalis assemblages.33 Recently, a database containing DNA sequence and epidemiological
data has been constructed55 to store information on human and animal isolates, which together comprises
about 4000 sequences from four genetic loci markers (18S rDNA, tpi, bg, and gdh). Analysis of the multilocus genotyping (MLG) data for assemblage A revealed the existence of three subgroups: subgroup
AI, which is predominantly found in livestock but is also present in humans and is likely to represent the
zoonotic type; subgroup AII, which is mostly found in humans (only a single cat isolate had an MLG also
found in human) and thus represents the human type; and subgroup AIII, which is essentially restricted to
wild animals, particularly wild ruminants, and is clearly restricted to animals (it has never been found in
humans).56 Therefore, when inferred using genetic data, zoonotic transmission of assemblage A appears
to be a rare event, but the lack of epidemiological links between the isolates investigated suggests that
caution should be used in generalizing these results. Indeed, a few studies have combined a designed
sampling strategy with molecular genotyping of the isolates. A study in a remote tea-growing community
in northern India, where humans and dogs live in close contact, has provided epidemiological evidence
supporting the role of dogs in the transmission to humans, albeit the molecular data were less convincing.57 Another study in temple communities in Bangkok (Thailand) has provided more epidemiological
and molecular evidence to support the role of dogs as reservoirs for human infection.58
Thus, the role of animals in the transmission of G. duodenalis assemblage A needs to be studied in the
context of subassemblages. Zoonotic transmission of assemblage AI appears to be minimal in developed
countries, but the situation may be different in endemic regions, for example, in localized foci where
humans and animal live promiscuously. Transmission of assemblage AII mostly occurs between humans
and may also take place from humans to animals. Zoonotic transmission of assemblage AII seems possible, but how often this happens is uncertain.
As reported earlier for assemblage A, the analysis of protein polymorphisms has been at the basis of
the recognition of genetic heterogeneity in G. duodenalis assemblage B isolates, with the recognition of
two subgroups BIII and BIV. Assemblage B isolates are more heterogeneous than assemblage A isolates,
to the point that assemblage B isolates were all different when compared at 27 genetic loci.6 In 2003,
Monis and colleagues10 analyzed 10 isolates at 23 genetic loci and further supported the existence of four
subgroups (BI, BII, BIII, and BIV). As was the case for assemblage A, human isolates appear to form two
clusters (subgroups BIII and BIV), whereas animal isolates (monkeys and a dog) belonged to subgroups
BI and BII. The single human isolate (BAH12) in subgroup BIII was, however, closer to subgroups BI
and BII. Therefore, zoonotic potential appears to be minimal, if any. More recent studies based on DNA
sequence analyses have suggested that the validity of these subgroups is questionable. Indeed, phylogenetic analysis of multiple genes did not support the clustering of human and animal isolates into strongly
supported, distinct clusters.23,35
Biology of Foodborne Parasites
As was the case for assemblage A, DNA sequence analyses revealed extensive polymorphism at all loci
investigated among assemblage B isolates. The rapid accumulation of sequence data allowed confirmation
that the level of genetic variation is higher in assemblage B compared to assemblage A.33 At first glance,
a higher genetic variability should facilitate molecular epidemiological studies, allowing better resolution in addressing research questions such as zoonotic transmission or the clinical significance of genotypes. However, direct sequencing of PCR products revealed the presence of “mixed positions” or “double
peaks,” that is, of two overlapping signals at specific positions in the sequencing profiles.59,60 This phenomenon was absent or very rare in sequences obtained from homologous loci of assemblage A. Indeed,
screening of assemblage B sequences in the Giardia database revealed that 21% of 1151 sequences (from
four different loci) contained “mixed” profiles, as compared to 5% in 1250 assemblage A sequences.55
Formally, these sequencing profiles can be generated by the genuine presence of different parasites (mixed
infections) and/or to allelic sequence heterogeneity (ASH) of a single population of parasites, but it is difficult to distinguish between the two situations when direct sequencing of PCR products is used. An important and elegant contribution on this issue was recently published.61 The authors used micromanipulation
to isolate single trophozoites from the assemblage B (GS isolate) of G. duodenalis, as well as single cysts
from human patients. They showed, through sequence analysis of PCR products at the tpi locus, that ASH
is present in single trophozoites from the GS lineage. Furthermore, ASH was demonstrated at the level of
single Giardia cysts of assemblage B from clinical samples by molecular analysis at the “beta-giardin”
and tpi loci. Additionally, alignment of sequence data from several different cysts that originated from the
same patient yielded different sequence patterns, thus suggesting the presence of multiple subassemblage
infections in congruence with ASH within the same patient.61 Therefore, genotyping of assemblage B isolates appears problematic and it will be necessary to identify genomic regions where ASH is low to design
new markers that can form the basis of a new genotyping scheme.
10.6 Pathogenesis and Clinical Features
Ingestion of as few as 10 cysts is considered sufficient to initiate a Giardia infection.1 The clinical spectrum of the infection is wide and ranges from the absence of overt symptoms (asymptomatic giardiasis)
to acute or chronic giardiasis.62 Symptoms appear 1 or 2 weeks after ingestion of cysts and persist for
3–4 days; the main symptom is diarrhea, but flatulence, epigastric cramps, nausea, vomiting, weight
loss, itch, and urticaria can also be observed.62 All these symptoms have been correlated to dysfunctions
of the small intestine, including a decrease in the surface area of brush border, atrophy of microvillus
and villus, enterocyte immaturity, deficiencies of luminal enzymes, and malabsorption of electrolytes,
fats, D-xylose, lactose, vitamin A, and vitamin B12.1 Malabsorption of nutrients and electrolytes creates
an osmotic gradient that draws water into the small intestinal lumen and that results in small intestinal
distension, rapid peristalsis, and, finally, diarrhea.
Exacerbation of symptoms has been observed in infants and children, especially in developing countries, where giardiasis has been associated with the failure-to-thrive syndrome, retarded growth and
development, poor cognitive function, and detrimental effects on nutritional status.63 A systematic
­follow-up of patients after a large outbreak that occurred in Bergen, Norway, has put into focus the fact
that giardiasis may be responsible for long-term consequences and sequelae in a naïve population. About
10% of infected individuals had persistent symptoms, with mean disease duration of 7 months, and at
least 5% of them developed persistent postinfectious fatigue syndrome.52 Furthermore, case reports and
epidemiological studies have associated giardiasis with the development of irritable bowel syndrome,
allergies, and reactive arthritis.64 Even if Giardia does not colonize extraintestinal sites, the infection can
lead to immunomediated disorders that can affect organs other than the intestine.65
The mechanisms that contribute to disease are only partially understood but are clearly multifactorial.65 For example, in the case of human asymptomatic infections, it is unclear if patients are infected
with a “nonpathogenic” assemblage/genotype of Giardia or if the host is able to keep parasite numbers
at a subclinical level without achieving its complete clearance. Certainly, both host and parasite factors
contribute to the pathogenesis of giardiasis and the existence of genotype/assemblage-specific pathogenic mechanisms is conceivable65 (Figure 10.3).
Barrier disruption
Immune cells
FIGURE 10.3 Graphical overview of the mechanism and effectors involved in the pathogenesis of giardiasis. (a)
Uninfected enterocytes. (b) Enterocytes infected with Giardia trophozoites. Interaction with the parasite induces enterocytes’ apoptosis and disruption of tight junctions and of intestinal barrier and can lead to malabsorption (unabsorbed
molecules are indicated by triangles). Infection also triggers recruitment of host immune cells (DC, dendritic cells; B,
lymphocytes B; T, lymphocytes T) and release of specific antibodies (indicated by a Y).
The first event necessary for the establishment of the infection depends on the high mobility and
strong attachment of the trophozoite to enterocytes in the upper small intestine. This is accomplished by
the ventral adhesive disk and by the flagella and avoids parasite elimination by peristalsis. The importance of the attachment process is clearly demonstrated by the reduced capacity of adhesion-deficient
Giardia clones to establish infection in Mongolian gerbils.66 The trophozoite must survive in the very
hostile environment of the small intestine, where it is readily exposed to proteases, lipases, bile salts, and
products of the host’s immune response. Giardia is the only gut-dwelling eukaryotic organism known
to possess a mechanism of antigenic variation.67 The surface of trophozoites is covered by a dense coat
of VSP, and a single VPS is normally dominating in a population of parasites. The on–off switching in
the expression of genes encoding VSP (approximately 270 genes in the WB strain) allows the parasite
to replace the expressed VSP during infection, possibly to escape the host immune reaction. Indeed,
primary infection with transgenic trophozoites simultaneously expressing multiple VSP or immunization with purified VSP from the transgenic cells protects gerbils from infection.68 Notably, the VSP
repertoire of different assemblages is totally different and no identical VSP are found in the genome
of the three strains sequenced to date.12,13 The involvement of secreted proteins with toxin-like activities has been hypothesized,62 but no giardial toxin has been identified thus far. However, four parasite
proteins, arginine deiminase (ADI), ornithine carbamoyltransferase (OCT), α-enolase, and elongation
factor 1-α, have been found in the supernatant of trophozoites cocultured with human intestinal epithelial
cells.69 In particular, the secretion of ADI and OCT seems to be involved in the inhibition of the innate
immune response applied by the host interfering with synthesis of nitric oxide (NO), which is cytostatic
to Giardia parasites, by subtracting L-arginine necessary for NO synthesis by the host epithelial cell
nitric oxide synthetase.70 Cysteine proteinase activities are also released upon contact with host cells and
may play a role in the adhesion of Giardia to epithelial cells, but the identity of these proteases has not
yet been established.71
A key component in the pathogenesis of Giardia infection is damage to enterocyte epithelium. An
increase in the rate of enterocyte apoptosis has been observed in in vitro experiments,72 shortly after colonization of the small intestine, and has been reported also in patients with chronic giardiasis.73 Apoptosis
appears to be mediated by activation of proapoptotic caspase-3 and caspase-9, increased expression
of proapoptotic Bax, decreased expression of antiapoptotic Bcl-2, and induced proteolytic cleavage of
Biology of Foodborne Parasites
poly(ADP-ribose) polymerase (PARP). Genes associated with apoptosis are indeed upregulated in cells
exposed to products from Giardia WBC6. Following the induction of apoptosis, Giardia trophozoites
can also induce alterations in the enterocyte tight junctions by the breakdown/relocalization of proteins
associated with these structures. Indeed, F-actin, zonula-occludens-1 (ZO-1), claudin-1, and α-actinin
are relocated from the cell periphery to the cytosol.65 Inhibition of caspase-3 prevents relocalization of
F-actin and ZO-1, suggesting a direct cause–effect relationship between Giardia-induced apoptosis and
small intestinal barrier function.72 The breakdown of epithelial barrier allows for macromolecules and
electrolytes to pass into the submucosa, bypassing normal uptake by epithelial cells. The paracellular
flow of nutrients and electrolytes can contribute to nutrient malabsorption by reducing electrochemical
gradients needed for proper uptake and can cause inflammation in some individuals, probably through
activation of innate immune effectors like macrophages.74 Finally, excess ion secretion was recently
demonstrated ex vivo in biopsies from patients suffering chronic giardiasis, suggesting that this may also
contribute to pathology.73
Very limited information is available on host factors directly involved in Giardia infection, albeit host
responses clearly play a key role in the pathogenesis of giardiasis. All the parasite-secreted proteins, the
VSP, and the major disk proteins are recognized by sera of Giardia-infected humans and by sera from
infected mice, indicating their importance in antibody-mediated Giardia immunity. Evidence obtained
from Giardia infection in animal models has suggested that IgA antibodies contribute to protective
immunity against giardiasis.74 However, resolution of the infection occurs even in mice that are unable
to produce antibodies.75 A relevant role of T-cell responses for the control of Giardia infections has been
demonstrated. In fact, T-cell-deficient mice develop chronic giardiasis75 and patients with common variable immunodeficiency (CVID) and Bruton’s X-linked agammaglobulinemia (XLA) have been associated with a predisposition to chronic giardiasis.29 Furthermore, a decreased mucosal surface area for the
absorption of nutrients, electrolytes, and water has been associated with a CD8+ lymphocyte-dependent
shortening of microvilli in the murine model.76
10.7 Treatment and Prevention
Although giardiasis is usually a self-limiting disease, treatment of confirmed cases is necessary to cure
symptoms and shorten the duration of acute infection, therefore reducing the risk of postinfectious complications and limiting environmental contamination with infectious cysts. Despite its clinical importance, no vaccines for human use have been approved thus far. A vaccine based on a total lysate of
Giardia trophozoites is available to treat cats and dogs (GiardiaVax©); the vaccine can reduce symptoms
and cyst shedding.77
Six classes of drugs are approved for treatment of giardiasis, including 5-nitroimidazole and benzimidazole derivatives, quinacrine, furazolidone, paromomycin, and nitazoxanide.78 Treatment is usually the same
for both immunocompetent and immunosuppressed patients. 5-nitroimidazoles are imidazole derivatives
that contain a nitro group, NO2, in position 5. MTZ (commercially Flagyl) is the best known of this class of
drugs. MTZ was demonstrated to be effective against G. duodenalis as early as in 1962 and is considered
the first-line antigiardial agent with a cure rate of 80%–95%.79 The recommended dose of MTZ is 500 mg
every 8 h for 7–10 days or 2 g for 3–5 days. MTZ is effective on trophozoites but has no effect on the viability
of cysts.80 MTZ and other nitroimidazoles are prodrugs and form active metabolites when the nitro moiety
is reduced in the cytoplasm of trophozoites.78 Purified Giardia pyruvate/ferredoxin oxidoreductase (PFOR)
and ferredoxin are involved in the reductive activation of MTZ.81 MTZ acts as an electron sink, due to
its lower redox potential compared to ferredoxin, and draws electrons away from ferredoxin. This reduction occurs only under intense reducing conditions, because oxygen can compete with MTZ as an electron
acceptor; this explains the selective toxicity of MTZ for anaerobic cells. Other enzymes such as a 2[4Fe-4S]
ferredoxin–nitroreductase fusion protein, an oxygen-insensitive nitroreductase (GlNR1), and thioredoxin
reductase are involved in MTZ activation in G. duodenalis.78,82 The formation of covalent adducts with
proteins and nonprotein thiols, as well as the interaction with DNA, has been demonstrated and is thought
to be part of the toxicity mechanism of nitroimidazole drugs in Giardia.82 Despite its efficacy, a number of
side effects have been reported for MTZ, including gastrointestinal upset, headache, nausea, leukopenia,
and a metallic taste in the mouth. Vomiting, flushing, headache, and gastrointestinal pain reactions may
occur after consumption of alcohol. Carcinogenicity and mutagenicity of MTZ have also been reported in
studies in rats and mice but have not been clinically proven.40,78 An approximately 10%–20% prevalence of
clinical MTZ-resistant cases of giardiasis has been reported,62 with recurrence rates of up to 90%. Various
resistant clones have been isolated from human patients, and MTZ-resistant clones have been induced in
different laboratories to study resistance mechanisms. Apparent resistance to MTZ and other nitroimidazole
compounds has been thereby linked to a decrease in reducing power, hence limiting prodrug activation.
Reduced expression or activity of PFOR and ferredoxin, thioredoxin reductase and suppressed flavin reduction, and nitroreductase GlNR1 has been described as molecular effectors involved in resistance mechanisms to nitro compounds.82 Laboratory-induced drug resistance has been also linked to antigenic variation
and altered expression of genes involved in signal transduction and network formation,17,83 suggesting that
resistance could be part of an adaptive mechanism to respond to environmental stress factors, including drug
pressure. MTZ resistance has been also associated with a significant fitness cost, as shown by the attenuated
infectivity of resistant strains in animal models; this results from an impaired ability of trophozoites to attach
that is linked to a reduced glucose metabolism.84 Alternative, long-acting 5-nitroimidazoles are available,
optimized for single daily dose therapies and with less side effects compared to MTZ. Tinidazole (commercially as Fasigyn) has been proved 100% successful in a single oral dosage of 2 g or 50 mg/kg.85 Secnidazole
(commercially Flagentyl) administrated in a single oral dose of 2 mg achieves clinical/parasitological cure
rates of 80%–100%, whereas ornidazole (commercially Tiberal) is generally administrated as a single dose
but for several days and was 100% effective in a trial conducted in Malaysia.78
In the case of treatment failure of first-line drugs (i.e., MTZ and albendazole), other nitro drugs can be
used as valid alternatives. The synthetic nitrofuran furazolidone (Furoxone) was introduced in 1960 for
the treatment of giardiasis. A cure rate between 80% and 96% has been reported for 7–10-day treatment
courses.79 It is administered as four doses per day in adults (100 mg/dose) and children (1.5 mg/kg). In
10% of patients, gastrointestinal disturbance, nausea, vomiting, diarrhea, malaise, itch, urticaria, hypersensitivity, and hemolysis in glucose-6-phosphate dehydrogenase-deficient patients have been reported.86
Despite lower efficacy, furazolidone is more efficient than MTZ in reducing cyst production in vitro. As
other nitro compounds, furazolidone is activated by reduction in the trophozoites whenever a reduced
flavin is present.87 Nitazoxanide (Alinia or Annita) is a synthetic nitrothiazolyl-salicylamide derivative
discovered in the 1980s as an antihelminthic88 and then found to be efficacious against Giardia and
other luminal parasites. This drug is approved in the United States for the treatment of cryptosporidiosis and giardiasis.89 A dose of 500 mg of nitazoxanide twice a day for 3 days is recommended, with an
85% clinical cure and 71%–80% parasite eradication observed in patients.88 The drug is well tolerated,
with few gastrointestinal side effects.78 Nitazoxanide is rapidly deacetylated in vivo to tizoxanide, which
has an equal effectiveness against the parasite. Even if the mode of action of nitazoxanide has not yet
fully elucidated, it has been shown that alterations in the ventral disk and surface membrane occur in
trophozoites treated in vitro with nitazoxanide, suggesting that the drug could induce defects in both
parasite attachment and nutrients’ absorption.90 Benzimidazoles, such as the benzimidazole carbamate
compound albendazole, fenbendazole, and mebendazole, exert a good antigiardial along with antihelminthic activity.91 Treatment of patients with albendazole was less effective than treatment with MTZ,
with a 62%–95% average efficacy of albendazole compared to 97% for MTZ.92 However, a recently
performed meta-analysis has shown that albendazole, when administered as a single dose of 400 mg/day
for 5 days, is as effective as MTZ in treatment of giardiasis.93 Side effects for benzimidazoles are similar
to those of MTZ but less intense.93 In parasitic nematodes, benzimidazoles block microtubule formation
by direct binding to the high-affinity binding site of β-tubulin monomer and abolishing dimerization
with α-tubulin.94 Similarly, in Giardia, benzimidazoles affect the rate and the amount of microtubules
assembling, and in trophozoites, this results in the disassociation of microtubule aggregates from the
adhesive disk and their dispersion in the cytoplasm.95 As for MTZ, cyst formation and viability are not
affected by the presence of benzimidazoles.95 Resistance to albendazole can be readily induced in vitro
with observable changes in the cytoskeletal structure without mutations of β-tubulin.83
In cases of seriously refractory giardiasis, the acridine yellow quinacrine can be used.96 Introduced
in the 1930s as an antimalarial, quinacrine was the first effective antigiardial drug, with a reported cure
rate of 92%–95%.97 However, quinacrine can cause serious side effects including dizziness, headache,
Biology of Foodborne Parasites
vomiting, psychotic response, reproductive tract cancers, development of abnormal uterine lesions, ectopic pregnancy, prolonged amenorrhea, and fetal exposure. Quinacrine treatment of Giardia trophozoites
is associated with a decrease in oxygen consumption, likely by inhibition of NADH oxidase,80 whereas
resistance is associated with an increased exclusion of the drug.98 Treatment of cysts with quinacrine
results in a reduced excystation rate.
During pregnancy, paromomycin is considered the safest drug to treat giardiasis, especially in the
first trimester. Paromomycin sulfate is a broad-spectrum aminoglycoside antibiotic isolated in 1950s
from Streptomyces rimosus var. paromomycinus.99 With a 60%–70% efficacy against giardiasis, paromomycin is poorly absorbed after oral administration and is mainly passed with feces without being
metabolized; consequently, no drug seems to reach the fetus.79,100 Nausea, increased gastrointestinal
motility, abdominal pain, and diarrhea are the most common side effects of paromomycin and, like other
aminoglycosides, systemic absorption of this antibiotic may cause ototoxicity and n­ ephrotoxicity.100
Paromomycin is an inhibitor of protein synthesis that, interacting with the prokaryotic ribosome’s 16S
and 23S rRNAs, affects the normal dissociation and recycling of the ribosomal subunits.101
Prevention is mainly based on good hygienic practices, such as frequent hand washing with soap
and by avoiding contact with the feces of infected persons. Travelers in countries where giardiasis is
endemic should use bottled water and avoid the consumption of raw fruits and vegetables and exposure
to untreated water (lakes, rivers, and streams).
1. Adam, R.D., Biology of Giardia lamblia. Clin. Microbiol. Rev., 14, 447, 2001.
2. Palm, D. et al., Developmental changes in the adhesive disk during Giardia differentiation. Mol. Biochem.
Parasitol., 141, 199, 2005.
3. Marti, M. et al., An ancestral secretory apparatus in the protozoan parasite Giardia intestinalis. J. Biol.
Chem., 278, 24837, 2003.
4. Jedelský, P.L. et al., The minimal proteome in the reduced mitochondrion of the parasitic protist Giardia
intestinalis. PLoS One, 6, e17285, 2011.
5. Rivero, M.R. et al., Receptor-mediated endocytosis and trafficking between endosomal-lysosomal vacuoles in Giardia lamblia. Parasitol. Res., 112, 1813, 2013.
6. Thompson, R.C.A. and Monis, P.T., Giardia- from genome to proteome. Adv. Parasitol., 78, 57, 2012.
7. Monis, P.T., Cacciò, S.M., and Thompson, R.C.A., Variation in Giardia: Towards a taxonomic revision
of the genus. Trends Parasitol., 25, 93, 2009.
8. Farthing, M.J., Varon, S.R., and Keusch, G.T., Mammalian bile promotes growth of Giardia lamblia in
axenic culture. Trans. R. Soc. Trop. Med. Hyg., 77, 467, 1983.
9. Monis, P.T. et al., Genetic diversity within the morphological species Giardia intestinalis and its relationship to host origin. Infect. Gen. Evol., 3, 29, 2003.
10. Monis, P.T. et al., Molecular systematics of the parasitic protozoan Giardia intestinalis. Mol. Biol. Evol.,
16, 1135, 1999.
11. Cacciò, S.M. and Ryan, U., Molecular epidemiology of giardiasis. Mol. Biochem. Parasitol., 160, 75,
12. Franzen, O. et al., Draft genome sequencing of Giardia intestinalis assemblage B isolate GS: Is human
giardiasis caused by two different species? PLoS Pathog., 5, e1000560, 2009.
13. Jerlstrom-Hultqvist, J. et al., Genome analysis and comparative genomics of a Giardia intestinalis
assemblage E isolate. BMC Genomics, 11, 543, 2010.
14. Svärd, S.G., Hagblom, P., and Palm, J.E., Giardia lamblia a model organism for eukaryotic cell differentiation. FEMS Microbiol. Lett., 218, 3, 2003.
15. Argüello-Garciá, R., Bazán-Tejeda, M.L., and Ortega-Pierres, G., Encystation commitment in Giardia
duodenalis: A long and winding road. Parasite, 16, 247, 2009.
16. Birkeland, S.R. et al., Transcriptome analyses of the Giardia lamblia life cycle. Mol. Biochem. Parasitol.,
174, 62, 2010.
17. Mueller, J. et al., Identification of differentially expressed genes in a Giardia lamblia WB C6 clone resistant to nitazoxanide and metronidazole. J. Antimicrob. Chemother., 62, 72, 2008.
18. Ringqvist, E. et al., Transcriptional changes in Giardia during host-parasite interactions. Int. J. Parasitol.,
41, 277, 2010.
19. Morf, L. et al., The transcriptional response to encystation stimuli in Giardia lamblia is restricted to a
small set of genes. Eukaryot. Cell, 9, 1566, 2010.
20. Morrison, H.G. et al., Genomic minimalism in the early diverging intestinal parasite Giardia lamblia.
Science, 317, 1921, 2007.
21. Yu, L.Z., Birky, C.W., and Adam, R.D., The two nuclei of Giardia each have complete copies of the
genome and are partitioned equationally at cytokinesis. Eukaryot. Cell, 1, 191, 2002.
22. Cooper, M.A. et al., Population genetics provides evidence for recombination in Giardia. Curr. Biol., 17,
1984, 2007.
23. Lasek-Nesselquist, E. et al., Genetic exchange within and between assemblages of Giardia duodenalis.
J. Eukaryot. Microbiol., 56, 504, 2009.
24. Ramesh, M.A., Malik, S.B., and Logsdon, J.M. Jr., A phylogenomic inventory of meiotic genes; evidence
for sex in Giardia and an early eukaryotic origin of meiosis. Curr. Biol., 15, 185, 2009.
25. Poxleitner, M.K. et al., Evidence for karyogamy and exchange of genetic material in the binucleate intestinal parasite Giardia intestinalis. Science, 319, 1530, 2008.
26. Cacciò, S.M. and Sprong, H., Giardia duodenalis: Genetic recombination and its implications for taxonomy and molecular epidemiology. Exp. Parasitol., 124, 107, 2010.
27. Smith, H.V. and Mank, T.G., Diagnosis of human giardiasis. In: Lujan, H.D. and Svard, S. (Eds.),
Giardia: A Model Organism. Springer, Wien, Austria, pp. 353–378, 2011.
28. Josko, D., Updates in immunoassays: Parasitology. Clin. Lab. Sci., 25, 185, 2012.
29. Faubert, G., Immune response to Giardia duodenalis. Clin. Microbiol. Rev., 13, 35, 2000.
30. Palm, J.E. et al., Identification of immunoreactive proteins during acute human giardiasis. J. Infect. Dis.,
12, 1849, 2003.
31. Ten Hoeve, R. et al., Detection of diarrhea-causing protozoa in general practice patients in the Netherlands
by multiplex real-time PCR. Clin. Microbiol. Infect., 13, 1001, 2007.
32. Lalle, M. et al., Genetic heterogeneity at the β-giardin locus among human and animal isolates of Giardia
duodenalis and identification of potentially zoonotic sub-genotypes. Int. J. Parasitol., 35, 207, 2005.
33. Wielinga, C.M. and Thompson, R.C., Comparative evaluation of Giardia duodenalis sequence data.
Parasitology, 134, 1795, 2007.
34. Sulaiman, I.M. et al., Triosephosphate isomerase gene characterization and potential zoonotic transmission of Giardia duodenalis. Emerg. Infect. Dis., 9, 1444, 2003.
35. Cacciò, S.M. et al., Multilocus genotyping of Giardia duodenalis reveals striking differences between
assemblages A and B. Int. J. Parasitol., 38, 1523, 2008.
36. Lebbad, M. et al., From mouse to moose: Multilocus genotyping of Giardia isolates from various animal
species. Vet. Parasitol., 168, 231, 2009.
37. Caccio, S.M. and Sprong, H., Epidemiology of giardiasis in humans. In: Lujan, H.D. and Svard, S.
(Eds.), Giardia: A Model Organism. Springer, Wien, Austria, pp. 16–28, 2011.
38. Savioli, L., Smith, H., and Thompson, R.C.A., Giardia and Cryptosporidium join the ‘Neglected Diseases
Initiative’. Trends Parasitol., 22, 203, 2006.
39. Feng, Y. and Xiao, L., Zoonotic potential and molecular epidemiology of Giardia species and giardiasis.
Clin. Microbiol. Rev., 24, 110, 2011.
40. Escobedo, A.A. et al., Giardiasis: The ever-present threat of a neglected disease. Infect. Disord. Drug
Targets, 10, 329, 2010.
41. Stark, D. et al., Clinical significance of enteric protozoa in the immunosuppressed human population.
Clin. Microbiol. Rev., 22, 634, 2009.
42. Yoder, J.S. et al., Giardiasis surveillance—United States, 2009–2010. MMWR Surveill. Summ., 61, 13,
43. Hoque, E. et al., A descriptive epidemiology of giardiasis in New Zealand and gaps in surveillance data.
NZ J. Med., 117, U1149, 2004.
44. Chute, C.G., Smith, R.P., and Baron, J.A., Risk factors for endemic giardiasis. Am. J. Public Health, 77,
585, 1987.
45. Stuart, J.M. et al., Risk factors for sporadic giardiasis: A case-control study in Southwestern England.
Emerg. Infect. Dis., 9, 229, 2003.
Biology of Foodborne Parasites
46. Espelage, W. et al., Prevalence and clinical correlations of genetic subtypes of Giardia lamblia in an
urban setting. BMC Public Health, 10, 41, 2010.
47. Faustini, A. et al., The impact of the Catholic Jubilee in 2000 on infectious diseases. A case-control study
of giardiasis, Rome, Italy 2000–2001. Epidemiol. Infect., 134, 649, 2006.
48. Hoque, M.E. et al., Nappy handling and risk of giardiasis. Lancet, 357, 1017, 2001.
49. Robertson, L.J. and Lim, Y.A.L., Waterborne and environmentally-borne giardiasis. In: Lujan, H.D. and
Svard, S. (Eds.), Giardia a Model Organism. Springer, Wien, Austria, pp. 29–69, 2011.
50. Baldursson, S. and Karanis, P., Waterborne transmission of protozoan parasites: Review of worldwide
outbreaks–An update 2004–2010. Water Res., 45, 6603, 2011.
51. Nygard, K. et al., A large community outbreak of waterborne giardiasis-delayed detection in a nonendemic urban area. BMC Public Health, 6, 141, 2006.
52. Naess, H. et al., Chronic fatigue syndrome after Giardia enteritis: Clinical characteristics, disability and
long-term sickness absence. BMC Gastroenterol., 12, 13, 2012.
53. Smith, H.V. et al., Cryptosporidium and Giardia as foodborne zoonoses. Vet. Parasitol., 149, 29, 2007.
54. Cook, N. et al., Development of a method for detection of Giardia duodenalis cysts on lettuce and for
simultaneous analysis of salad products for the presence of Giardia cysts and Cryptosporidium oocysts.
Appl. Environ. Microbiol., 73, 7388, 2007.
55. Sprong, H., Cacciò, S.M., and van der Giessen, J., Identification of zoonotic genotypes of Giardia duodenalis. PLOS Neg. Trop. Dis., 3, e558, 2009.
56. Lalle, M. et al., A novel Giardia duodenalis assemblage A subtype in fallow deer. J. Parasitol., 93, 426,
57. Traub, R.J. et al., Epidemiological and molecular evidence support the zoonotic transmission of Giardia
among humans and dogs living in the same community. Parasitology, 128, 53, 2004.
58. Traub, R.J. et al., Transmission cycles of Giardia duodenalis in dogs and humans in temple communities
in Bangkok—A critical evaluation of its prevalence using three diagnostic tests in the field in the absence
of a gold standard. Acta Trop., 111, 125, 2009.
59. Gelanew, T. et al., Molecular characterization of human isolates of Giardia duodenalis from Ethiopia.
Acta Trop., 102, 92, 2007.
60. Lebbad, M., et al., Dominance of Giardia assemblage B in Leon, Nicaragua. Acta Trop., 106, 44, 2008.
61. Ankarklev, J., Svärd, S.G., and Lebbad, M., Allelic sequence heterozygosity in single Giardia parasites.
BMC Microbiol., 12, 65, 2012.
62. Farthing, M.J., The molecular pathogenesis of giardiasis. J. Ped. Gastroent. Nutr., 24, 79, 1997.
63. Berkman, D.S. et al., Effects of stunting, diarrhoeal disease, and parasitic infection during infancy on
cognition in late childhood: A follow-up study. Lancet, 59, 564, 2002.
64. Wensaas, K.A. et al., Irritable bowel syndrome and chronic fatigue 3 years after acute giardiasis: Historic
cohort study. Gut, 61, 214, 2012.
65. Cotton, J.A., Beatty, J.K., and Buret, A.G., Host parasite interactions and pathophysiology in Giardia
infections. Int. J. Parasitol., 41, 925, 2011.
66. Hernandez-Sanchez, J. et al., Giardia duodenalis: Adhesion-deficient clones have reduced ability to
establish infection in Mongolian gerbils. Exp. Parasitol., 119, 364, 2008.
67. Carranza, P.G. and Lujan, H.D., New insights regarding the biology of Giardia lamblia. Microbes Infect.,
12, 71, 2010.
68. Rivero, F.D. et al., Disruption of antigenic variation is crucial for effective parasite vaccine. Nat. Med.,
16, 551, 2010.
69. Skarin, H. et al., Elongation factor 1-alpha is released into the culture medium during growth of Giardia
intestinalis trophozoites. Exp. Parasitol., 127, 804, 2011.
70. Rópolo, A.S. and Touz, M.C., A lesson in survival, by Giardia lamblia. Scientific World J., 10, 2019, 2010.
71. Rodríguez-Fuentes, G.B. et al., Giardia duodenalis: Analysis of secreted proteases upon trophozoiteepithelial cell interaction in vitro. Mem. Inst. Oswaldo Cruz., 101, 693, 2006.
72. Chin, A.C. et al., Strain-dependent induction of enterocyte apoptosis by Giardia lamblia disrupts epithelial barrier function in a caspase-3-dependent manner. Infect. Immun., 70, 3673, 2002.
73. Troeger, H. et al., Effect of chronic Giardia lamblia infection on epithelial transport and barrier function
in human duodenum. Gut, 56, 328, 2007.
74. Solaymani-Mohammadi, S. and Singer, S., Giardia duodenalis: The double-edged sword of immune
responses in giardiasis. Exp. Parasitol., 126, 292, 2010.
75. Singer, S.M. and Nash, T.E., T-cell-dependent control of acute Giardia lamblia infections in mice. Infect.
Immun., 68, 170, 2000.
76. Scott, K.G., Yu, L.C., and Buret, A.G., Role of CD8+ and CD4+ T lymphocytes in jejunal mucosal injury
during murine giardiasis. Infect. Immun., 72, 3536, 2004.
77. Olson, M.E., Ceri, H., and Morck, D.W., Giardia vaccination. Parasitol. Today, 16, 213, 2000.
78. Lalle, M., Giardiasis in the post genomic era: Treatment, drug resistance and novel therapeutic perspectives. Infect. Disord. Drug Targets, 10, 283, 2010.
79. Gardner, T.B. and Hill, D.R., Treatment of giardiasis. Clin. Microbiol. Rev., 14, 114, 2011.
80. Paget, T.A. et al., Respiration in the cyst and trophozoite forms of Giardia muris. J. Gen. Microbiol., 135,
145, 1989.
81. Upcroft, P. and Upcroft, J.A., Drug targets and mechanisms of resistance in the anaerobic protozoa. Clin.
Microbiol. Rev., 14, 150, 2001.
82. Leitsch, D. et al., Pyruvate:ferredoxin oxidoreductase and thioredoxin reductase are involved in 5-nitroimidazole activation while flavin metabolism is linked to 5-nitroimidazole resistance in Giardia lamblia.
J. Antimicrob. Chemother., 66, 1756, 2011.
83. Argüello-García, R. et al., In vitro resistance to 5-nitroimidazoles and benzimidazoles in Giardia duodenalis: Variability and variation in gene expression. Infect. Genet. Evol., 9, 1057, 2009.
84. Tejman-Yarden, N. et al., Impaired parasite attachment as fitness cost of metronidazole resistance in
Giardia lamblia. Antimicrob. Agents Chemother., 55, 4643, 2011.
85. Fung, H.B. and Doan, T.L., Tinidazole: A nitroimidazole antiprotozoal agent. Clin. Ther., 27, 1859, 2005.
86. Ali, V. and Nozaki, T., Current therapeutics, their problems, and sulfur-containing-amino-acid metabolism as a novel target against infections by “amitochondriate” protozoan parasites. Clin. Microbiol. Rev.,
20, 164, 2007.
87. Leitsch, D. et al., Trichomonas vaginalis: Metronidazole and other nitroimidazole drugs are reduced by
the flavin enzyme thioredoxin reductase and disrupt the cellular redox system. Implications for nitroimidazole toxicity and resistance. Mol. Microbiol., 72, 518, 2009.
88. Rossignol, J.F., Ayoub, A., and Ayers, M.S., Treatment of diarrhea caused by Giardia intestinalis and
Entamoeba histolytica/dispar: A double blind placebo controlled study of nitazoxanide. J. Infect. Dis.,
184, 381, 2001.
89. Fox, L.M. and Saravolatz, L.D., Nitazoxanide: A new thiazolide antiparasitic agent. Clin. Infect. Dis., 40,
1173, 2005.
90. Muller, J. et al., In vitro effects of thiazolides on Giardia lamblia WB clone C6 cultured axenically and
in coculture with Caco2 cells. Antimicrob. Agents Chemother., 50, 162, 2006.
91. Morgan, U.M., Reynoldson, J.A., and Thompson, R.C.A., Activities of several benzimidazoles and tubulin inhibitors against Giardia spp. in vitro. Antimicrob. Agents Chemother., 37, 328, 1993.
92. Hall, A. and Nahar, Q., Albendazole as a treatment for infections with Giardia duodenalis in children in
Bangladesh. Trans. R. Soc. Trop. Med. Hyg., 87, 84, 1993.
93. Granados, C.E. et al., Drugs for treating giardiasis. Cochrane Database Syst. Rev., 12, CD007787, 2012.
94. Kwa, M.S. et al., β-Tubulin genes from the parasitic nematode Haemonchus contortus modulate drug
resistance in Caenorhabditis elegans. J. Mol. Biol., 246, 500, 1995.
95. Hausen, M.A. et al., Giardia lamblia: A report of drug effects under cell differentiation. Parasitol. Res.,
105, 789, 2009.
96. Mørch, K. et al., Treatment-ladder and genetic characterisation of parasites in refractory giardiasis after
an outbreak in Norway. J. Infect., 56, 268, 2008.
97. Wolfe, M.S. and Handler, R.P., Quinacrine for treatment of giardiasis. J. Travel Med., 5, 228, 1998.
98. Upcroft, P., Multiple drug resistance in the pathogenic protozoa. Acta Trop., 56, 195, 1994.
99. Davidson, R.N., den Boer, M., and Ritmeijer, K., Paromomycin. Trans. R. Soc. Trop. Med. Hyg., 103,
653, 2009.
100. Mineno, T. and Avery, M.A., Giardiasis: Recent progress in chemotherapy and drug development. Curr.
Pharm. Res., 9, 841, 2003.
101. Scheunemann, A.E. et al., Binding of aminoglycoside antibiotics to helix 69 of 23S rRNA. Nucleic Acids
Res., 8, 3094, 2010.
Benjamin Rosenthal
11.1 Introduction................................................................................................................................... 195
11.2 Biology, Genetics, and Genomics................................................................................................. 196
11.3 Diagnosis and Typing.................................................................................................................... 198
11.4 Epidemiology and Molecular Epidemiology................................................................................ 199
11.5 Pathogenesis and Clinical Features.............................................................................................. 201
11.6 Treatment and Prevention............................................................................................................. 203
11.7 Conclusions................................................................................................................................... 203
References............................................................................................................................................... 204
11.1 Introduction
Species of Sarcocystis and their most immediate relatives are ascribed to the subfamily Sarcocystidae.
They form tissue cysts in various tissues of their intermediate hosts, which can persist for prolonged
periods in a metabolically quiescent, developmentally arrested phase. These encysted stages are called
sarcocysts because they are predominantly found in muscle tissue of the intermediate host. To complete
the life cycle, all species of Sarcocystis require a second host (also termed the final or definitive host).
This host must be a carnivore that ingests the intramuscular cyst (with its infectious contents) and supports, in its intestinal tract, the parasite’s sexual development.1,2 Other parasites within this tissue-cystforming group include genera Besnoitia, Hammondia, Neospora, and Toxoplasma. These, too, have life
cycles that depend on predator–prey relationships.
Of the myriad vertebrate intermediate hosts infected by species of Sarcocystis, cattle, sheep, pigs,
poultry, water buffalo, and goats have been studied most carefully due to their economic importance.
Nonetheless, species of Sarcocystis have been found in every tetrapod group (including reptiles, amphibians, mammals, and birds). Although species of Sarcocystis have not yet been found in fish, they may
have evolved from parasites of fish.3
Among species of Sarcocystis that parasitize livestock, two are known to cause human foodborne
intestinal infection; people can contract intestinal sarcocystosis by consuming beef or pork infected
with Sarcocystis hominis or Sarcocystis suihominis, respectively (Figure 11.1). Exposure to human feces
enables those livestock to contract infections. Other species of Sarcocystis occur in cattle and swine
and complete their sexual development in feline or canine definitive hosts, instead of in human beings.
Consuming other raw meats may expose people to other forms of Sarcocystis, but no others have been
established as zoonotic. As discussed in greater detail in Sections 11.4 and 11.5, people can also serve
as intermediate hosts of Sarcocystis, but our understanding of the sources of these exposures remains
in its infancy.
Biology of Foodborne Parasites
Definitive host
Infected meat
eaten by humans
Cyst in muscle cells
Intermediate Gametes
Bovine and swine
ingested by
cow and pig
FIGURE 11.1 Life cycle of S. hominis and S. suihominis. Ingesting muscle cysts in either beef or pork results in GI infection, culminating in the excretion of sporulated oocysts.
11.2 Biology, Genetics, and Genomics
The two, well-defined cycles of human intestinal sarcocystosis result from eating raw or undercooked
beef or pork (Figure 11.1). Such meat may be infected with parasitic tissue cysts (sarcocysts). Each sarcocyst originates from a haploid progenitor (metrocyte) that undergoes many rounds of asexual division
within the host cell; in about 2 months, the mature and infectious sarcocyst will be packed with many
bradyzoites, the small, crescent-shaped form that defines the terminally differentiated state in the intermediate host (Figure 11.2). These sarcocysts occur in many muscle types, especially the diaphragm,
tongue, and esophagus. Some species (especially Sarcocystis neurona, as discussed in Section 11.5) have
a predilection for neural tissue.
When ingested by a carnivore, the previously sedentary bradyzoites become motile, exit the sarcocyst,
and penetrate the lamina propria of the intestine. Sexual differentiation then produces female “macrogamonts” and male “microgametocytes.” Their fusion produces a transient diploid phase, followed by meiosis
and formation of an oocyst containing eight haploid progenies. About 2 weeks later, oocysts exit the lamina
propria and enter the lumen of the gut, whereupon they are passed in the feces. A mature oocyst comprises
two sporocysts, each of which contains four sporozoites (Figure 11.3). The thin oocyst wall may or may not
be visible surrounding a pair of sporocysts; frequently, the oocyst wall will rupture prior to examination,
FIGURE 11.2 Mature sarcocyst of S. hominis. Numerous quiescent bradyzoites are surrounded by a cyst wall characterized by villar protrusions within a muscle cell. (Stained with Toluidine blue.)
FIGURE 11.3 Progressive maturation of oocysts derived from the lamina propria. Before sporulation commences (left),
the oocysts appear filled with unorganized granules. Partially sporulated oocysts (top, middle) divide these into two discrete aggregations. Final maturation (right) results in the appearance of two sporocysts, each containing four sporozoites
and a residual body. Mature sporocysts may appear as pairs (as shown). From fecal preparations, single sporocysts may be
recovered owing to fracture of the oocyst wall.
freeing each unpaired sporocyst (each containing four sporozoites) in the feces of the carnivore. Those
excreted sporocysts then can infect cattle or swine upon ingestion of contaminated water, food, or soil.
When livestock contract infection, the parasites do not remain confined to the gastrointestinal (GI)
tract, do not undergo sexual maturation, and are not excreted to the environment. Instead, each sporozoite undergoes asexual division, dissemination to various tissues and organs via the vascular system,
Biology of Foodborne Parasites
and develops into a long-lived tissue cyst. Further development awaits consumption by predation or
scavenging. A key galactose lectin that mediates host cell invasion survives the proteolytic environment of the GI tract of intermediate and definitive hosts by concealing two dozen hydrogen bonds,
responsible for dimerization, in a protected, highly hydrophobic pocket.4
Thus, the course and duration of infection differ markedly in definitive and intermediate hosts. In
definitive hosts (such as people in the case of S. hominis and S. suihominis), the parasite induces a fleeting bout of GI disease. In intermediate hosts (true for cattle and swine in the case of these two parasite
species), the parasite establishes chronic (indeed, lifelong) infections. As is true for any omnivore, people
may serve as either the definitive or intermediate host, resulting in acute or chronic infection.
Only rarely has it been suggested that the same host serves as both the intermediate and definitive
hosts for a given parasite species. Certain cannibalistic lizards, inhabiting islands lacking other carnivores, have been proposed to serve as both intermediate and definitive hosts of a parasite related to
Sarcocystis gallotiae.5 Likewise, rats have been identified as excreting sporocysts and also harboring tissue cysts for Sarcocystis cymruensis.6 Because cats were also shown to support the sexual development
of parasites derived from these rats, some skepticism seems warranted; it is possible that distinct parasite
types complete sexual development in cats and rats.
Although sporocysts typically engender new infections only after being excreted by definitive hosts,
unexcreted sporocysts might pose a threat to the health of predators of definitive hosts. This has been
postulated to explain fatal cardiac, pulmonary, and cerebral infections in eagles whose prey included
opossums (Didelphis virginiana) harboring, in their digestive tracts, sporocysts of S. neurona.7
Transmission can be extraordinarily efficient. The predation and carrion feeding practiced by fishers
(Martes pennanti), for example, result in almost universal exposure.8 Examples of Sarcocystis abound in
mammals, birds, amphibians, and reptiles, exploiting myriad predator–prey cycles. Only a few transmission cycles involving human hosts have been well characterized, but others may occur. Indeed, human
exposure varies according to food habits and hygiene, and scientific investigation will likely establish
new sources of human infection (as discussed in Section 11.7).
With the exception of a diploid period in the gut of the definitive host, species of Sarcocystis persist as
haploid cells. Oocysts contain two sporocysts, each of which contains four haploid sporozoites. Haploidy
is maintained throughout the prolonged infection in intermediate hosts, sustained by mitotic divisions
(giving rise to mobile merozoites), and then within the developing sarcocyst. The sarcocyst, after many
rounds of such replication, will encompass a large number of progeny derived through this strictly asexual division of haploid cells.
The genetics of Sarcocystis populations have begun to be studied. As might be expected for an organism that engages in prolonged intervals of asexual reproduction, and that can derive both male and
female gametes from the same haploid progenitor lineage (enabling “selfing”), certain clonal lineages
have been identified as responsible for given outbreaks.9,10 Broader sampling of endemic populations has
identified myriad multilocus genotypes, as might be expected for an entity that routinely undergoes a
sexual stage.11–13
S. neurona has been subjected to additional study of its cell biology, and expressed sequence libraries have provided information concerning the major metabolic pathways elaborated in these exclusively
intracellular parasites. Sequential expression of distinct surface antigens has been documented in S. neurona.14 More should soon be learned from the inclusion of S. neurona in ongoing comparative genome
sequencing efforts devoted to Toxoplasma gondii. However, the extent of differences among the genomes
or transcriptomes of different species in the genus remains to be determined.
11.3 Diagnosis and Typing
To diagnose human intestinal sarcocystosis, sporocysts must first be recovered from the stool; diagnosing muscular sarcocystosis requires the biopsy of muscle tissue. In either case, PCR amplification of
diagnostic loci aids efforts to confirm the presence of Sarcocystis spp. and is required to differentially
diagnose the causative agent. Those tools will be spelled out in greater detail in Section 11.4, after a brief
overview of the history of diagnosis and typing.
Microscopy greatly advanced the diagnosis of species of Sarcocystis but cannot, by itself, serve to
differentiate among excreted forms (sporocysts) owing to extensive intraspecific variation in their shape
and size and owing to overlapping size distributions among species. Greater diagnostic variation characterizes the appearance of mature muscle cysts; however, their appearance and size change as they
mature. Light microscopy enables sarcocysts to be subdivided according to their overall size (some
are microscopic, and others much longer) and suffices to categorize those with thick or thin sarcocyst
walls. Transmission electron microscopy (TEM) provides further means to differentiate among sarcocysts based on the pattern of invaginations in the sarcocyst wall at the interface of the parasite and its
host cell.15 A nomenclature of sarcocyst wall types has proved fairly robust to variations in cyst maturity
and histological preparation.
Molecular diagnosis has played an increasingly important role in differentiating among species and
strains, in part owing to the fact that experimental infections have only rarely fulfilled “Koch’s postulates,” establishing the capacity of a given species of Sarcocystis to cycle among a given pair of definitive and intermediate hosts. Though Sarcocystis muris was long ago shown to be capable of cycling
between cats and mice, natural infections in feral cats were only recently confirmed.16 Without molecular diagnostic methods, the sporocysts in these cats might have been mistaken for those of T. gondii.
A cycle involving rats and king rat snakes was recently completed experimentally, a study that also ruled
mice out as suitable intermediate hosts.17 Another study recently succeeded in transmitting Sarcocystis
gracilis from roe deer to each of two kinds of foxes.18 It is difficult to raise livestock under controlled
environments and is even more difficult to raise wildlife in captivity with defined, parasite-free diets.
Experimental infections have established transmission from swine to people (S. suihominis), from cattle
to people (S. hominis), from cattle to dogs (S. cruzi), and from a variety of intermediate hosts to opossums (S. neurona).
Genetic signatures have, therefore, contributed vital information necessary to indirectly establish
chains of transmission. For example, foxes and arctic foxes had been established as predators responsible
for transmitting certain parasites to moose (Alces alces).19 Other sarcocysts in moose were found to be
genetically very distinct, suggesting that they originated from other definitive hosts. Observing large
numbers of corvid birds scavenging on moose carcasses led some to suspect these as definitive hosts,
a suspicion subsequently confirmed by sequencing sporocysts derived from corvid feces that matched
the tissue cysts in the moose.20 The list of hosts from which S. neurona has been reported is growing
(detailed in Section 11.4); although the taxonomic status of an assemblage of parasites attributed to
Sarcocystis falcatula remains uncertain (it may be a single taxon, very closely related to S. neurona, or
an assemblage of remarkably similar taxa), finding these parasites in several families of birds came as
some surprise.7
11.4 Epidemiology and Molecular Epidemiology
Recent progress in molecular epidemiology is exemplified in studies of S. neurona transmission. This
agent initially garnered attention as the agent of a neurological disease in horses, equine protozoal
myeloencephalitis (EPM). Histology first established this condition as infectious in origin, and sequencing the 18S ribosomal DNA (rDNA) provided clues as to the type of organism that should be suspected
as its cause. Serendipity revealed genetically indistinguishable organisms in the feces of opossums (D.
­virginiana).21 This led to the successful demonstration that horses exposed to sporocysts shed by opossums would contract disease, though it remained unclear whether horses played a natural role in transmitting the infection or whether horses were merely aberrant (clinically symptomatic) hosts.
The search was on to identify other hosts that naturally harbored or could be experimentally shown
to be capable of harboring this agent. The first completion of the life cycle was accomplished in raccoons (Procyon lotor), and genetic evidence was offered as a means to validate its identity. Since then,
molecular evidence has established the occurrence of tissue cysts of S. neurona in a variety of naturally
infected hosts, including raccoons, cats, and armadillos, providing an understanding of disease transmission that could never have been accomplished by exclusive reliance on experimental infections.22
Indeed, opossums might never have been tested for their capacity to serve as definitive hosts; although
Biology of Foodborne Parasites
their feces do contaminate pastures, opossums do not routinely feed on horsemeat. Had efforts at establishing the main route of transmission been predicated on a central role for horses, the source of this
equine disease might have remained elusive.
Sequencing 18S rDNA has provided the best basis to differentiate among parasite isolates, determine
their relationships, and link the stages occurring in tissues to the stages occurring in feces. Differentiating
among closely related taxa has benefitted from the inclusion of other genes, including randomly amplified
genomic fragments.23,24 Recently, progress has been reported from successful attempts to amplify >1000
bases of cytochrome oxidase 1 encoded in the mitochondrial genome.25 Alone, and especially in combination, these markers provide an effective means to discriminate among species of Sarcocystis and to
reconstruct their evolutionary relationships. In so doing, they provide a basis to erect and test hypotheses
concerning transmission routes. These procedures are used by research groups in North America, Europe,
Asia, the Middle East, and South America. They remain in the domain of veterinary research but have not
become incorporated into routine medical or veterinary clinical practice. Variation in the sequence of 18S
rDNA among various parasite types induces differences in the temperature at which double-stranded amplicons denature to single strands. Such differences have been explored as a basis for accomplishing differential diagnosis using methods that do not require the PCR product to be sequenced, saving time and money.26
Finding sarcocysts in the tissues of a new intermediate host was once deemed grounds for naming a
new species; it was not then understood that an intermediate host might harbor the cysts of more than
one parasite species. Electron microscopy later increased the ability to discriminate among species of
Sarcocystis, including among those occurring in the same host. For example, a recent study used microscopy to determine that roe deer can harbor four distinct types of Sarcocystis.27 Cross transmission and
molecular evidence have demonstrated that closely related host species can be infected by the same
species of Sarcocystis. Red deer (Cervus elaphus) and reindeer (Rangifer tarandus), for example, can
be infected by the same species of Sarcocystis.28 The same is true for certain parasites that infect water
buffalo (Bubalus bubalis) and cattle (Bos taurus).29
Serological assays are available for testing exposure to certain species of Sarcocystis. The standard
ELISA was demonstrated to have only poor sensitivity when used on live animals.30 Improvements to
serology for S. neurona have been reported using a polyvalent assay that overcomes variable expression
of distinct surface antigens.31
Those species of Sarcocystis that infect ruminant intermediate hosts comprise an evolutionary clade
that is subdivided according to those using canine and feline definitive hosts.32 The phylogeny of parasites does not always recapitulate relationships among definitive hosts; overlapping feeding habits appear
to encourage host switches.33
Molecular tools have been instrumental in establishing that transmission can spill far beyond the
bounds of the natural cycle, at times causing serious disease in accidental hosts. For example, marked
declines in southern sea otters (Enhydra lutris) have been attributed to exposure to S. neurona.34 Episodes
of heavy rainfall precede, by 30–60 days, outbreaks among these marine mammals that elevate the risk
of fatal sarcocystosis to 12 times the risk of death from other causes.35 Thus, even marine mammals
inhabiting the coastal margin of islands at some distance from the sources of infection are vulnerable
to exposure, disease, and even death. The ability of Sarcocystis spp. to become established in atypical
settings, potentially compromising the health of protected and endangered species, is further exemplified
by their occurrence in an isolated population of dogs in the Galapagos Islands.36
In general, zoonotic infections span a spectrum from those that only poorly infect human beings (and
lack the capacity to be transmitted among people) to those that thrive on, and even depend on, human
hosts. Most successful human pathogens were contracted either from nonhuman primates or from our
livestock.37 In keeping with this general pattern, the species of Sarcocystis for which humans serve as
the definitive host derive from livestock species (swine, cattle); we should not be surprised to learn of
additional examples (i.e., in poultry, goats, sheep) from which people acquire infection and for which
humans serve as the source of infection. Likewise, we should suspect other primates as possible sources
of human sarcocystosis.
Evidence is emerging that other primates may, indeed, be the natural intermediate hosts that serve as a
reservoir for parasites capable of causing human muscular sarcocystosis. Some years ago, a parasite was
recognized in the macaque and designated as Sarcocystis nesbitti. Sarcocysts derived from that macaque
were recently subjected to molecular analysis, and the ensuing phylogenetic analysis placed this parasite
among a clade that uses carnivorous snakes as definitive hosts.38 The possibility that a predatory snake
may have been the source of sporocysts has increased, given recent reports that Malaysian snakes harbor
an assemblage of Sarcocystis types, including S. nesbitti.39 Furthermore, a parasite indistinguishable by
18S rDNA from S. nesbitti has recently been diagnosed in human infections acquired on Pangkor Island,
Malaysia.40 This finding not only substantiates the hypothesis that snakes may have been the source but
also affirms the notion that human beings are especially susceptible to those infections adapted to nonhuman primates. As a practical matter, such a finding underscores how important it is to safeguard our
drinking water from contamination with the feces of carnivores.
11.5 Pathogenesis and Clinical Features
As emphasized earlier, sarcocystosis is a tale of two infections: intestinal sarcocystosis for the definitive
host and muscular sarcocystosis for the intermediate host.
A few, well-documented cases of experimental human infection provide the basis for our understanding of intestinal sarcocystosis ensuing from ingestion of sarcocysts in raw pork or beef.41 Symptoms have
varied among people, and according to the infectious dose, but GI distress typically commences within
6–36 h; nausea, distension, and abdominal pain have been reported. After an interval of about 2 weeks,
and lasting for only a period of weeks, sporocysts appear in the stool. Thus, the intestinal sarcocystosis
that people experience after consuming infected pork or beef is limited in scope and duration. It has
not yet been established whether a person’s genetic makeup, nutritional status, or immunological status
markedly affects this outcome or whether ingesting sarcocysts from other kinds of meat induces clinically distinct outcomes.
Less information is available concerning the pathogenesis of human muscular sarcocystosis, although
recent outbreaks in Southeast Asia are shedding new light on that subject. Surveys of human biopsy have,
for some years, indicated that sarcocysts may be prevalent in certain human populations. Most notably,
a sample of tongues derived from postmortem samples indicated prevalent infections (~20%) in Thailand,
although the identity of the causative agent (or agents) remains unknown. A high population prevalence of
antibodies to Sarcocystis spp. has also been reported from Malaysia.42,43
A 1999 report documented the symptoms experienced by five members of a U.S. Military team
stationed in Malaysia who contracted muscular sarcocystosis after eating contaminated meat.44 More
clinical data are now being amassed from recent outbreaks of human sarcocystosis among travelers to
Tioman and Pangkor Islands, Malaysia. Fever, muscle pain, elevated creatine kinase, and eosinophilia
have been typical.40,45–47
A report concentrating on five such individuals47 described headache, fever, and pain in the joints,
muscles, back, and extremities. Watery diarrhea and hives were reported from one individual, who later
developed tachycardia, palpitations, and a recurring rash. Two patients experience pain that “moved
from the arms to the legs and then to the back from day to day” and in the thighs, calves, upper arms,
back, and (in one patient) the tongue. Normal electrocardiograms were observed in all but one patient,
who was diagnosed with an incomplete right bundle branch block and few ventricular premature beats.
Eosinophilia and elevated creatine kinase levels were observed in all cases. More than 100 individuals
have been identified as part of an ongoing outbreak on Tioman Island, and the clinical and epidemiological data that will ensue from a concerted scientific effort will surely reveal additional information
about the sources of infection, the course of illness, and the methods that prove generally effective in
treating afflicted individuals.46 Interestingly, the survey of this broader group has indicated a delayed
increase in creatine kinase and identified two initially asymptomatic cases (a child and an adult who
subsequently developed swollen skeletal and cardiac muscles severe enough to warrant hospitalization).
People infected with this form of sarcocystosis have generally experienced two phases of clinical symptoms, corresponding to a febrile acute stage of infection, followed by a longer-lasting phase of inflammation and muscle pain (Doug Esposito, personal communication). Another Malaysian outbreak of human
sarcocystosis was recently reported from the island of Pangkor, in which the most prominent symptoms
among 89 cases were myalgia, headache, cough, and inflammatory edema.40
Biology of Foodborne Parasites
As valuable as these outbreak investigations are to understanding the etiology of human muscular
sarcocystosis, it must be borne in mind that the clinical experience of travelers (experiencing new exposures as adults) may be very different from persons who encounter routine exposures beginning early
in life. The reportedly high prevalence of infection in Thailand, but the absence of infection in Ireland,
underscores the fact that the clinical picture may vary markedly from place to place.48,49 It is likely that
human beings are exposed to a variety of sporocysts (shed by any number of carnivorous hosts) and that
many of these are incapable of completing development of mature sarcocysts in us (resulting in abortive,
clinically inapparent infections). Others may occasionally induce aberrant infections (akin to S. neurona
in a horse) resulting in a wide range of clinical outcomes. However, human burial processes generally
protect cadavers from routine scavenging by other animals, and human beings are seldom any longer at
risk of predation.
Pathology in other animals is clearly variable. S. cruzi is fairly ubiquitous in cattle; molecular data indicate that this parasite has disseminated wherever dogs and cattle occur.50,51 Primary infection induces fever
and myalgia, but bovine sarcocystosis generally poses no significant threat to the health and well-being
of cattle. A recent survey of lesions was performed in condemned cattle carcasses in Europe. In ~80% of
the cases where sarcocysts could be characterized (using a combination of laser-capture microdissection
and sequencing), S. hominis was diagnosed. However, immune responses leading to lesions significant
enough to warrant carcass condemnation were also attributed to S. cruzi, S. hirsuta, and an unidentified
species,52 underscoring the variable clinical outcomes that can ensue even from infections with generally
avirulent parasites. In another study, injecting antigens sufficed to induce bovine eosinophilic myositis.53
The same cannot be said for S. canis, a parasite first identified in dogs and subsequently diagnosed as the
cause of severe liver infections in Steller sea lions, black bears, and polar bears.54–58 It is unclear how commonly such liver disease results from infection. Whether or not they are typical clinical outcomes, these
cases illustrate a marked pathology associated with certain forms of sarcocystosis in intermediate hosts.
So severe are the sequelae in certain types of rats infected with Sarcocystis singaporensis that it is
used as a rodenticide.59,60 This parasite uses pythons as its definitive host. Presumably, other rodents suffer less from such infections, but certain parasites in certain hosts frequently result in death.
The example of S. neurona nicely illustrates the variety of responses that hosts can mount to infection with a given species of Sarcocystis. Recall that S. neurona was first recognized as a cause of EPM,
resulting in debilitating ataxia in horses. Horses may or may not play a role in sustaining cycles of transmission of this agent. Horses that are seropositive for S. neurona do not always exhibit clinical signs and
do not consistently diminish in their antibody titers after experimental treatment, but severe disease in a
range of other hosts has been attributed to infections with S. neurona.61,62
Another neurological syndrome in pigeons, pigeon protozoal myeloencephalitis, has recently gained
attention as an emerging disease and a model for studying immune disregulation.63 Here, immune
responses seem to be thwarted during the early stages of infection, but delayed hypersensitivity may
later induce debilitating neurological lesions. The recent identification of this parasite in European sparrow hawks (Accipiter nisus) and wood pigeons (Columba palumbus) suggests that domestic pigeons
(Columba livia domestica), which suffer such marked disease, may represent accidental hosts (as has
been suggested for horses and S. neurona).63
Similarly, Sarcocystis tenella was recently implicated as the cause of severe pneumonia in several
sheep, resulting in the death of three individuals, in spite of the fact that asymptomatic muscle infections are routine.64 Swine are commonly infected with Sarcocystis meisheriana, a parasite acquired
by ingesting sporocysts shed by cats. Although such infections are typically subclinical, this parasite
was diagnosed as the cause of death for a breeding boar that experienced multifocal degeneration and
necrosis of myocardial fibers with interstitial edema, severe multifocal nonsuppurative myocarditis and
hepatitis, and nonsuppurative interstitial nephritis.65 Marine mammals infected with multiple species
(Sarcocystis and T. gondii) suffered especially grave symptoms.66 Whether this indicates interactions
leading to worse pathology, or disproportionate vulnerability of certain animals, remains uncertain.
A fatal, systemic infection of S. neurona in a skunk (Mephistis mephistis) may have been exacerbated by
simultaneous infection with canine distemper virus.67
Quite clearly, the symptoms caused by chronic infections with Sarcocystis vary considerably among
species of parasites and among hosts encountering infection with a given parasite.
11.6 Treatment and Prevention
Clinical data on human sarcocystosis remain too limited to substantiate any particular course of treatment. Three of four individuals suspected of contracting sarcocystosis on Tioman Island, Malaysia,
showed clinical improvement after two daily treatments of albendazole (400 mg each) for 2 weeks followed by prednisone for 1 week (80, 40, and 20 mg/day in decreasing dosage) or 20 mg/day for 4 weeks.
The fourth patient experienced relapse and was subjected to a second round of treatment.47
Successful treatment of a California sea lion infected with S. neurona was reported using a 28-day
course of ponazuril (10 mg/kg [4.5 mg/lb], PO, q 24 h) and a 5-day treatment with prednisone (0.2 mg/kg
[0.09 mg/lb], PO, q 12 h).56 Decoquinate was also successful in treating one of two dogs suffering massive infections with an undetermined species of Sarcocystis.68 Decoquinate, a quinolone anticoccidial
approved for use in the prevention of intestinal coccidiosis in farm animals, was recently shown to kill
cultures of S. neurona at nanomolar concentrations, suggesting that it might bear investigation as a
treatment in vivo.69 Similar promise has been shown for nitazoxanide and new thiazolide/thiadiazolide
efficacy against S. neurona, in vitro.70
Adequately cooking meat represents the simplest way to prevent human exposure to either of the two
known forms of Sarcocystis for which we serve as definitive hosts (or for any other such parasite that has
not yet been characterized). Deep freezing can also achieve that goal. Pork cooked at 60°C for 20 min,
at 70° for 15 min, or at 100° for 5 min was rendered uninfectious. Freezing at −4°C for 48 h, or −20°C
for 24 h, was also efficacious.71 A recent report from Japan claims that freezing horsemeat at −20°C
for 48 h or more eliminated the toxicity of a protein attributed to Sarcocystis fayeri.72 Although such a
modality for pathogenesis has not been thoroughly studied, it has been observed that rabbits exposed to
bradyzoites suffered fatal hypersensitivity reactions (J.P. Dubey, personal communication) suggesting
that pathology may result from exposure to parasite proteins even when those parasites are nonviable.
Hygienic management of human wastes reduces exposure of our livestock to sporocysts, and therefore,
one would expect only limited transmission from people to livestock where wastewater is treated. In this
regard, it is somewhat surprising that a significant proportion of tested beef in Europe harbors infections
with S. hominis. Unless there are also nonhuman definitive hosts contributing to bovine exposure, this
fact indicates that cattle are routinely being exposed to raw human sewage. The same has not been found
in surveys of beef from the Americas; here, most cattle harbor sarcocysts of S. cruzi, contracted from dog
feces, and are seldom diagnosed with S. hominis.73,50
Prevention, as outlined in the two preceding paragraphs, would minimize transmission cycles that rely
on human hosts. How best to prevent human exposure to parasites that naturally cycle among nonhuman
hosts? Safety would increase with physical distance from such enzootic cycles, particularly those cycles
involving nonhuman primates. Moreover, clean sources of food and water would confer general protection.
In the outbreaks being subjected to ongoing investigation, there is reason to believe that nonpotable
water, or surface waters vulnerable to contamination with animal feces, may have infiltrated the sources
of water for drinking and food preparation. Any practice that increased human exposure to fecal contamination would represent a general risk to human health, an insight that dates to the very founding
of epidemiology when cholera outbreaks were mapped by John Snow to particular sources of water in
London. In the specific case of sporocysts, the risk would derive from the feces of carnivores.
11.7 Conclusions
We should not be surprised if, in coming years, the list of species in this diverse genus undergoes revision. Occasionally, it has been determined that the same parasite occurs in different (typically related)
host types. Electron microscopy and molecular diagnostics have also helped verify that a given host
may be infected with more than one species of Sarcocystis. Undoubtedly, many await scientific attention. For example, three new pathogens were recently encountered in a study of infected sea lions.74
The true diversity of the genus will not be known until every potential host has been assayed and each
identified parasite type has been unequivocally speciated. Such a concerted effort would undoubtedly
Biology of Foodborne Parasites
yield results, as contemporary diagnostic methodologies (pairing ultrastructural analysis of the sarcocyst
wall, sequencing, and phylogenetic analysis) have proven quite effective in clarifying the diversity and
relationships among these myriad parasite types.75,76 Indeed, a recent report suggests that one of the best
studied intermediate hosts (cattle) in one of the best studied regions (Europe) harbors a parasite whose
definitive host remains unknown, but which appears most prevalent on farms where people consume raw
meat and where sewage treatment is lacking.77 Thus, additional zoonotic parasites may await characterization in even the most conventional of food sources. Given the breadth of the human diet, and variation
in culinary practices, the actual zoonotic impact of human sarcocystosis may be far greater than what the
clinical literature has heretofore established.
1. Fayer, R., Sarcocystis: Development in cultured avian and mammalian cells, Science, 168, 1104, 1970.
2. Fayer, R., Gametogony of Sarcocystis sp. in cell culture, Science, 175, 65, 1972.
3. Molnar, K., Ostoros, G., Dunams-Morel, D., and Rosenthal, B.M., Eimeria that infect fish are diverse
and are related to, but distinct from, those that infect terrestrial vertebrates, Infect. Genet. Evol., 12, 1810,
4. Müller, J.J., Weiss, M.S., and Heinemann, U., PAN-modular structure of microneme protein SML-2
from the parasite Sarcocystis muris at 1.95 Å resolution and its complex with 1-thio-β-D-galactose, Acta
Crystallogr. D Biol. Crystallogr., 67, 936, 2011.
5. Harris, D.J., Maia, J.P., and Perera, A., Molecular survey of Apicomplexa in Podarcis wall lizards detects
Hepatozoon, Sarcocystis, and Eimeria species, J. Parasitol., 98, 592, 2012.
6. Hu, J.J. et al., Identification of Sarcocystis cymruensis in wild Rattus flavipectus and Rattus norvegicus
from Peoples Republic of China and its transmission to rats and cats, J. Parasitol., 97, 421, 2011.
7. Wunschmann, A. et al., Natural fatal Sarcocystis falcatula infections in free-ranging eagles in North
America, J. Vet. Diagn. Invest., 22, 282, 2010.
8. Larkin, J.L. et al., Prevalence to Toxoplasma gondii and Sarcocystis spp. in a reintroduced fisher (Martes
pennanti) population in Pennsylvania, J. Parasitol., 97, 425, 2011.
9. Wendte, J.M. et al., Self-mating in the definitive host potentiates clonal outbreaks of the apicomplexan
parasites Sarcocystis neurona and Toxoplasma gondii, PLoS Genet., 6, e1001261, 2010.
10. Wendte, J.M. et al., Limited genetic diversity among Sarcocystis neurona strains infecting southern sea
otters precludes distinction between marine and terrestrial isolates, Vet. Parasitol., 169, 37, 2010.
11. Monteiro, R.M. et al., Extensively variable surface antigens of Sarcocystis spp. infecting Brazilian marsupials in the genus Didelphis occur in myriad allelic combinations, suggesting sexual recombination has
aided their diversification, Vet. Parasitol., 196, 64, 2013.
12. Asmundsson, I.M., Dubey, J.P., and Rosenthal, B.M., A genetically diverse but distinct North American
population of Sarcocystis neurona includes an overrepresented clone described by 12 microsatellite
alleles, Infect. Genet. Evol., 6, 352, 2006.
13. Sundar, N. et al., Modest genetic differentiation among North American populations of Sarcocystis neurona may reflect expansion in its geographic range, Vet. Parasitol., 152, 8, 2008.
14. Gautam, A., Dubey, J.P., Saville, W.J., and Howe, D.K., The SnSAG merozoite surface antigens of
Sarcocystis neurona are expressed differentially during the bradyzoite and sporozoite life cycle stages,
Vet. Parasitol., 183, 37, 2011.
15. Dubey, J.P., Speer, C.A., and Fayer, R., Sarcocystis of Animal and Man, CRC Press, Boca Raton, FL,
16. Kappany, A. et al., Experimental transmission of Sarcocystis muris (Coccidia: Apicomplexa) from the
feces of a naturally infected feral cat (Felis catus) from Egypt, J. Parasitol., 95, 1036, 2013.
17. Hu, J.J., Meng, Y., Guo, Y.M., Liao, J.Y., and Song, J.L., Completion of the life cycle of Sarcocystis zuoi,
a parasite from the Norway rat, Rattus norvegicus, J. Parasitol., 98, 550, 2012.
18. Gjerde, B., Morphological and molecular characterization and phylogenetic placement of Sarcocystis
capreolicanis and Sarcocystis silva n. sp. from roe deer (Capreolus capreolus) in Norway, Parasitol.
Res., 110, 1225, 2012.
19. Dahlgren, S.S. and Gjerde, B., The red fox (Vulpes vulpes) and the arctic fox (Vulpes lagopus) are definitive
hosts of Sarcocystis alces and Sarcocystis hjorti from moose (Alces alces), Parasitology, 137, 1547, 2010.
20. Gjerde, B. and Dahlgren, S.S., Corvid birds (Corvidae) act as definitive hosts for Sarcocystis ovalis in
moose (Alces alces), Parasitol. Res., 107, 1445, 2010.
21. Fenger, C.K. et al., Identification of opossums (Didelphis virginiana) as the putative definitive host of
Sarcocystis neurona, J. Parasitol., 81, 916, 1995.
22. Dubey, J.P. et al., A review of Sarcocystis neurona and equine protozoal myeloencephalitis (EPM), Vet.
Parasitol., 95, 89, 2001.
23. Tanhauser, S.M. et al., Multiple DNA markers differentiate Sarcocystis neurona and Sarcocystis falcatula, J. Parasitol., 85, 221, 1999.
24. Rosenthal, B.M., Lindsay, D.S., and Dubey, J.P., Relationships among Sarcocystis species transmitted by
New World opossums (Didelphis spp.), Vet. Parasitol., 95, 133, 2001.
25. Gjerde, B., Phylogenetic relationships among Sarcocystis species in cervids, cattle and sheep inferred
from the mitochondrial cytochrome c oxidase subunit I gene, Int. J. Parasitol., 43, 579, 2013.
26. Lalonde, L.F. and Gajadhar, A.A., Detection and differentiation of coccidian oocysts by real-time PCR
and melting curve analysis, J. Parasitol., 97, 725, 2011.
27. Pérez-Creo, A. et al., Prevalence and identity of Sarcocystis spp. in roe deer (Capreolus capreolus) in
Spain: A morphological study, Res. Vet. Sci., 2013.
28. Dahlgren, S.S. and Gjerde, B., Molecular characterization of five Sarcocystis species in red deer (Cervus
elaphus), including Sarcocystis hjorti n. sp., reveals that these species are not intermediate host specific,
Parasitology, 137, 815, 2010.
29. Jehle, C. et al., Diagnosis of Sarcocystis spp. in cattle (Bos taurus) and water buffalo (Bubalus bubalis)
in Northern Vietnam, Vet. Parasitol., 166, 314, 2009.
30. Johnson, A.L., Burton, A.J., and Sweeney, R.W., Utility of 2 immunological tests for antemortem diagnosis of equine protozoal myeloencephalitis (Sarcocystis neurona Infection) in naturally occurring cases,
J. Vet. Intern. Med., 24, 1184, 2010.
31. Yeargan, M.R. and Howe, D.K., Improved detection of equine antibodies against Sarcocystis neurona
using polyvalent ELISAs based on the parasite SnSAG surface antigens, Vet. Parasitol., 176, 16, 2011.
32. Holmdahl, O.J., Morrison, D.A., Ellis, J.T., and Huong, L.T., Evolution of ruminant Sarcocystis
(Sporozoa) parasites based on small subunit rDNA sequences, Mol. Phylogenet. Evol., 11, 27, 1999.
33. Tome, B., Maia, J.P., and Harris, D.J., Molecular assessment of apicomplexan parasites in the snake
psammophis from North Africa: Do multiple parasite lineages reflect the final vertebrate host diet?
J. Parasitol., 99, 883, 2013.
34. Miller, M.A. et al., A protozoal-associated epizootic impacting marine wildlife: Mass-mortality of southern sea otters (Enhydra lutris nereis) due to Sarcocystis neurona infection, Vet. Parasitol., 172, 183, 2010.
35. Shapiro, K., Miller, M., and Mazet, J., Temporal association between land-based runoff events and
California sea otter (Enhydra lutris nereis) protozoal mortalities, J. Wildl. Dis., 48, 394, 2012.
36. Gingrich, E.N., Scorza, A.V., Clifford, E.L., Olea-Popelka, F.J., and Lappin, M.R., Intestinal parasites of
dogs on the Galapagos Islands, Vet. Parasitol., 169, 404, 2010.
37. Wolfe, N.D., Dunavan, C.P., and Diamond, J., Origins of major human infectious diseases, Nature, 447,
279, 2007.
38. Tian, M. et al., Phylogenetic analysis of Sarcocystis nesbitti (Coccidia: Sarcocystidae) suggests a snake
as its probable definitive host, Vet. Parasitol., 183, 373, 2012.
39. Lau, Y.L. et al., Genetic assemblage of Sarcocystis spp. in Malaysian snakes, Parasit. Vectors, 6, 257,
40. Abubakar, S. et al., Outbreak of human infection with Sarcocystis nesbitti, Malaysia, 2012, Emerg. Infect.
Dis., 19, 1989, 2013.
41. Fayer, R., Sarcocystis spp. in human infections, Clin Microbiol Rev., 17, 894, 2004.
42. Kan, S.P. and Pathmanathan, R., Review of sarcocystosis in Malaysia, Southeast Asian J. Trop. Med.
Public Health, 22(Suppl), 129, 1991.
43. Wong, K.T. and Pathmanathan, R., High prevalence of human skeletal muscle sarcocystosis in south-east
Asia, Trans. R. Soc. Trop. Med. Hyg., 86, 631, 1992.
44. Arness, M.K., Brown, J.D., Dubey, J.P., Neafie, R.C., and Granstrom, D.E., An outbreak of acute eosinophilic myositis attributed to human Sarcocystis parasitism, Am. J. Trop. Med. Hyg., 61, 548, 1999.
45. CDC, Notes from the field: Acute muscular sarcocystosis among returning travelers—Tioman Island,
Malaysia, 2011, MMWR, 61, 37, 2012.
Biology of Foodborne Parasites
46. Esposito, D.H., Freedman, D.O., Neumayr, A., and Parola, P., Ongoing outbreak of an acute muscular Sarcocystis-like illness among travellers returning from Tioman Island, Malaysia, 2011–2012, Euro.
Surveill., 17, 2012.
47. Tappe, D. et al., Initial patient cluster and first positive biopsy findings in an outbreak of acute muscular Sarcocystis-like infection in travelers returning from Tioman island, Peninsular Malaysia, in 2011,
J. Clin. Microbiol., 51, 725, 2013.
48. Pathmanathan, R. and Kan, S.P., Three cases of human Sarcocystis infection with a review of human
muscular sarcocystosis in Malaysia, Trop. Geogr. Med., 44, 102, 1992.
49. Wong, K.T., Leggett, P.F., and Heatley, M., Apparent absence of Sarcocystis infection in human tongue
and diaphragm in Northern Ireland, Trans. R. Soc. Trop. Med. Hyg., 87, 496, 1993.
50. Pritt, B. et al., Detection of Sarcocystis parasites in retail beef: A regional survey combining histological
and genetic detection methods, J. Food. Prot., 71, 2144, 2008.
51. Rosenthal, B.M., Dunams, D.B., and Pritt, B., Restricted genetic diversity in the ubiquitous cattle parasite, Sarcocystis Cruzi, Infect. Genet. Evol., 8, 588, 2008.
52. Vangeel, L. et al., Different Sarcocystis spp. are present in bovine eosinophilic myositis, Vet. Parasitol.,
197, 543, 2013.
53. Vangeel, L. et al., Intramuscular inoculation of cattle with Sarcocystis antigen results in focal eosinophilic myositis, Vet. Parasitol., 183, 224, 2012.
54. Allison, R. et al., Fatal hepatic sarcocystosis in a puppy with eosinophilia and eosinophilic peritoneal
effusion, Vet. Clin. Pathol., 35, 353, 2006.
55. Garner, H.M. et al., Fatal hepatic sarcocystosis in two polar bears (Ursus maritimus), J. Parasitol., 83,
523, 1997.
56. Carlson-Bremer, D.P., Gulland, F.M., Johnson, C.K., Colegrove, K.M., and Van Bonn, W.G., Diagnosis
and treatment of Sarcocystis neurona-induced myositis in a free-ranging California sea lion, J. Am. Vet.
Med. Assoc., 240, 324, 2012.
57. Welsh, T., Burek-Huntington, K., Savage, K., Rosenthal, B., and Dubey, J.P., Sarcocystis canis associated
hepatitis in a Steller sea lion (Eumetopias jubatus) from Alaska, J. Wildl. Dis., 50, 405, 2014.
58. Davies, J.L., Haldorson, G.J., Bradway, D.S., and Britton, A.P., Fatal hepatic sarcocystosis in a captive
black bear (Ursus americanus) associated with Sarcocystis canis-like infection, J. Vet. Diagn. Invest., 23,
379, 2011.
59. Jakel, T., Burgstaller, H., and Frank, W., Sarcocystis singaporensis: Studies on host specificity, pathogenicity, and potential use as a biocontrol agent of wild rats, J. Parasitol., 82, 280, 1996.
60. Jakel, T. et al., Biological control of rodents using Sarcocystis singaporensis, Int. J. Parasitol., 29, 1321,
61. Pusterla, N. et al., Evaluation of the kinetics of antibodies against Sarcocystis neurona in serum from
seropositive healthy horses without neurological deficits treated with ponazuril paste, Vet. Rec., 173, 249,
62. Britton, A.P., Dubey, J.P., and Rosenthal, B.M., Rhinitis and disseminated disease in a ferret (Mustela
putorius furo) naturally infected with Sarcocystis neurona, Vet. Parasitol., 169, 226, 2010.
63. Olias, P. et al., Modulation of the host Th1 immune response in pigeon protozoal encephalitis caused by
Sarcocystis calchasi, Vet. Res., 44, 10, 2013.
64. Schock, A. et al., Respiratory disease due to acute Sarcocystis tenella infection in sheep, Vet. Rec., 170,
571, 2012.
65. Caspari, K., Grimm, F., Kuhn, N., Caspari, N.C., and Basso, W., First report of naturally acquired clinical
sarcocystosis in a pig breeding stock, Vet. Parasitol., 177, 175, 2011.
66. Gibson, A.K. et al., Polyparasitism is associated with increased disease severity in Toxoplasma gondiiinfected marine sentinel species, PLoS Negl. Trop. Dis., 5, e1142, 2011.
67. Burcham, G.N., Ramos-Vara, J.A., and Vemulapalli, R., Systemic sarcocystosis in a striped skunk
(Mephitis mephitis), Vet. Pathol., 47, 560, 2010.
68. Sykes, J.E. et al., Severe myositis associated with Sarcocystis spp. infection in 2 dogs, J. Vet. Intern.
Med., 25, 1277, 2011.
69. Lindsay, D.S., Nazir, M.M., Maqbool, A., Ellison, S.P., and Strobl, J.S., Efficacy of decoquinate against
Sarcocystis neurona in cell cultures, Vet Parasitol., 196, 21, 2013.
70. Gargala, G. et al., In vitro efficacy of nitro- and halogeno-thiazolide/thiadiazolide derivatives against
Sarcocystis neurona, Vet. Parasitol., 162, 230, 2009.
71. Saleque, A., Juyal, P.D., and Bhatia, B.B., Effect of temperature on the infectivity of Sarcocystis miescheriana cysts in pork, Vet. Parasitol., 36, 343, 1990.
72. Harada, S. et al., Control of toxicity of Sarcocystis fayeri in horsemeat by freezing treatment and prevention of food poisoning caused by raw consumption of horsemeat, Shokuhin Eiseigaku Zasshi., 54, 198,
73. Moré, G. et al., Development of a multiplex real time PCR to differentiate Sarcocystis spp. affecting
cattle, Vet. Parasitol., 197, 85, 2013.
74. Colegrove, K.M. et al., Discovery of three novel coccidian parasites infecting California sea lions
(Zalophus californianus), with evidence of sexual replication and interspecies pathogenicity, J. Parasitol.,
97, 868, 2011.
75. Dubey, J.P. et al., Two new species of Sarcocystis (Apicomplexa: Sarcocystidae) infecting the wolverine
(Gulo gulo) from Nunavut, Canada, J. Parasitol., 96, 972, 2010.
76. Prakas, P., Kutkienė, L., Butkauskas, D., Sruoga, A., and Zalakevičius, M., Molecular and morphological
investigations of Sarcocystis corvusi sp. nov. from the jackdaw (Corvus monedula), Parasitol. Res., 112,
1163, 2013.
77. Domenis, L. et al., Detection of a morphogenetically novel Sarcocystis hominis-like in the context of a
prevalence study in semi-intensively bred cattle in Italy, Parasitol. Res., 109, 1677, 2011.
Toxoplasma gondii
Dolores E. Hill and Jitender P. Dubey
12.1 Introduction................................................................................................................................... 209
12.2 Morphology and Classification......................................................................................................210
12.3 Biology, Genetics, and Genomics..................................................................................................215
12.4 Diagnosis........................................................................................................................................215
12.5 Epidemiology.................................................................................................................................215
12.6 Pathogenesis and Clinical Features...............................................................................................216
12.7 Treatment and Prevention..............................................................................................................218
12.8 Future Directions and Trends........................................................................................................219
12.1 Introduction
Toxoplasma gondii is a coccidian parasite with an unusually wide range of intermediate hosts. Felids
serve as definitive hosts and produce the environmentally resistant oocyst stage. Toxoplasmosis is one
of the most common parasitic infections of man, though its prevalence varies widely from place to
place. It continues to be a significant public health problem in the United States, where 8%–22% of
people are infected; a similar prevalence is seen in the United Kingdom.1–5 In Central America, South
America, and continental Europe, estimates of infection range from 30% to 90%.2,6,7 Most infections in
humans are asymptomatic, but at times, the parasite can produce devastating disease. Infection may be
congenitally or postnatally acquired. In the United States, nationwide serological surveys demonstrated
that seroprevalence in people remained stable at 23% from 1990 until 1998,3 while recent surveys have
demonstrated a significant decrease in seroprevalence to 10.8% over the last decade.5 Similar decreases
in seroprevalence have been observed in many European countries.6
It is estimated that 1,075,242 persons are infected with T. gondii each year in the United States, and
approximately 2,839 persons develop symptomatic ocular disease annually.8 The cost of illness in the
United States caused by Toxoplasma has been estimated to be nearly three billion dollars and an 11,000
quality-adjusted life year (QALY) loss annually.9,10 Recent publications have linked suicide and schizophrenia to Toxoplasma infection.11,12
T. gondii also infects food animals, including sheep, goats, pigs, chickens, and many game animal species. Infected animals harbor tissue cysts, and human consumers can be infected by ingestion of these
cysts in raw or undercooked meat. Virtually all edible portions of an animal can harbor viable T. gondii
tissue cysts,13 and tissue cysts can survive in food animals for years. The relative contribution of foodborne
(meat) sources of Toxoplasma infection versus oocyst transmission of Toxoplasma to human infection
is unknown, and various studies have suggested widely disparate estimates of foodborne transmission.
Mead et al.14 suggested that T. gondii is one of the three pathogens (along with Salmonella and Listeria)
that account for >75% of all deaths due to foodborne disease in the United States. Roghmann et al.15 suggested that 50% of Toxoplasma infections in the United States could be ascribed to foodborne transmission. Scallan et al.16 estimated that Toxoplasma caused 8% of hospitalizations and 24% of deaths resulting
Biology of Foodborne Parasites
from foodborne illnesses. In contrast, Dubey et al.,17 in a nationwide survey of retail meats (beef, chicken,
and pork), found no viable Toxoplasma in any of 2094 beef or 2094 chicken samples and 7 positive pork
samples out of 2094 samples assayed, concluding that there was not enough viable Toxoplasma present
in retail meats to account for the level of Toxoplasma infection found in the U.S. population. Recent
studies18,19 have suggested that oocyst exposure is the predominate route of Toxoplasma transmission
in the United States. Despite the uncertainty of human infection source, Toxoplasma is recognized as a
foodborne risk. Animal infections with Toxoplasma, especially infections in non-meat-eating ruminants,
birds, and pigs raised in confinement, also likely result from environmental exposure to T. gondii oocysts.
Oocyst contamination of the environment is widespread as a result of fecal contamination of soil and
groundwater by the estimated 140 million domestic and feral cats in the United States, each of which can
deposit hundreds of millions of oocysts in feces during infection.6,20 Oocyst-contaminated runoff surface water entering the marine environment has resulted in devastating disease in endangered sea otters
off the west coast of the United States,21,22 and even wild herbivores have been shown to have very high
seroprevalence as a result of exposure to the environmentally resistant oocysts.23
12.2 Morphology and Classification
T. gondii belongs to phylum Apicomplexa (Levine, 1970), class Sporozoasida (Leukart, 1879), subclass
Coccidiasina (Leukart, 1879), order Eimeriorina (Leger, 1911), and family Toxoplasmatidae (Biocca,
1956). There is only one species of Toxoplasma, T. gondii. Coccidia in general have complicated life
cycles. Most coccidia are host specific and are transmitted via a fecal–oral route. Transmission of
T. ­gondii occurs via the fecal–oral route (Figure 12.1), as well as through consumption of infected meat,
and by transplacental transfer from mother to fetus.1,24
The name Toxoplasma (toxon = arc, plasma = form) is derived from the crescent shape of the
tachyzoite stage (Figure 12.2). There are three infectious stages of T. gondii: the tachyzoites (in groups)
(Figure 12.3a), the bradyzoites (in tissue cysts) (Figure 12.3b and c), and the sporozoites (in oocysts)
FIGURE 12.1 Life cycle of T. gondii.
Toxoplasma gondii
FIGURE 12.2 Tachyzoites of T. gondii. Bar = 10 µm. (a) Individual (small arrows), binucleate (large arrow), and divided
(arrowhead) tachyzoites. Impression smear of lung. Compare size with red blood cells and leukocytes. Giemsa stain.
(b) Tachyzoites in a group (large arrow) and in pairs (small arrows) in section of a mesenteric lymph node. Note organisms
are located in PVs, and some are dividing (arrowhead). Hematoxylin and eosin (H & E) stain.
(f )
FIGURE 12.3 Stages of T. gondii. Scale bar in a–d = 20 µm and in e–g = 10 µm. (a) Tachyzoites in impression smear
of a lung. Note crescent-shaped individual tachyzoites (arrows) and dividing tachyzoites (arrowheads) compared with the
size of host red blood cells and leukocytes. Giemsa stain. (b) Tissue cysts in section of muscle. The tissue cyst wall is very
thin (arrow) and encloses many tiny bradyzoites (arrowheads). H & E stain. (c) Tissue cyst separated from host tissue by
homogenization of infected brain. Note tissue cyst wall (arrow) and hundreds of bradyzoites (arrowheads). Unstained.
(d) Schizont (arrow) with several merozoites (arrowheads) separating from the main mass. Impression smear of infected cat
intestine. Giemsa stain. (e) A male gamete with two flagella (arrows). Impression smear of infected cat intestine. Giemsa
stain. (f) Unsporulated oocyst in fecal float of cat feces. Unstained. Note double-layered oocyst wall (arrow) enclosing a
central undivided mass. (g) Sporulated oocyst with a thin oocyst wall (large arrow), two sporocysts (arrowheads). Each
sporocyst has four sporozoites (small arrow), which are not in complete focus. Unstained.
Biology of Foodborne Parasites
1 μm
FIGURE 12.4 Transmission electron micrograph (TEM) of a tachyzoite of T. gondii in a mouse peritoneal exudate cell;
Am, amylopectin granule; Co, conoid; Dg, electron-dense granule; Fp, fingerlike projection of tachyzoite plasmalemma;
Go, Golgi complex; Hc, host cell cytoplasm; Im, inner-membrane complex; Mi, mitochondrion; Mn, microneme; Nu,
nucleus; Pl, plasmalemma; Pv, parasitophorous vacuole; Rh, rhoptry; Sm, subpellicular microtubule; Tv, tubulovesicular
membranes. Bar = 1 µm.
(Figure 12.3g). The tachyzoite is often crescent shaped and is approximately the size (2 × 6 µm) of
a red blood cell (Figure 12.4). The anterior end of the tachyzoite is pointed, and the posterior end
is round. It has a pellicle (outer covering), several organelles including subpellicular microtubules,
mitochondrium, smooth and rough endoplasmic reticulums, a Golgi apparatus, apicoplast, ribosomes,
a micropore, and a well-defined nucleus. The nucleus is usually situated toward the posterior end or
in the central area of the cell. The tachyzoite enters the host cell by active penetration of the host
cell membrane and can tilt, extend, and retract, as it searches for a host cell. After entering the host
cell, the tachyzoite becomes ovoid in shape and becomes surrounded by a parasitophorous vacuole
(PV; Figure 12.4). T. gondii in a PV is protected from host defense mechanisms. The tachyzoite multiplies asexually within the host cell by repeated divisions in which two progenies form within the
parent parasite, consuming it (Figure 12.5a through d). Tachyzoites continue to divide until the host
cell is filled with parasites.
After a few divisions, T. gondii forms tissue cysts that vary in size from 5 to 70 µm and remain intracellular (Figure 12.6a through f). The tissue cyst wall is elastic, thin (<0.5 µm), and may enclose hundreds of the crescent-shaped, slender T. gondii stage known as bradyzoites (Figure 12.7). The bradyzoites
are approximately 7 × 1.5 µm in size and differ structurally only slightly from tachyzoites. They have a
nucleus situated toward the posterior end, whereas the nucleus in tachyzoites is more centrally located.
Bradyzoites are more slender and less susceptible to destruction by proteolytic enzymes than tachyzoites. Although tissue cysts containing bradyzoites may develop in visceral organs, including lungs, liver,
and kidneys, they are more prevalent in muscular and neural tissues, including the brain (Figure 12.6a
through f), eye, skeletal, and cardiac muscle. Intact tissue cysts probably do not cause any harm and can
persist for the life of the host.
Toxoplasma gondii
FIGURE 12.5 T. gondii stages in in vitro and in vivo preparations. (a) Tachyzoites in culture of human foreskin fibroblast
cells. Giemsa stain. Bar = 25 µm. (b) Rosettes of tachyzoites in human foreskin fibroblasts. Immunohistochemical stain
with anti-tachyzoite-specific antibody. Smear. Bar = 10 µm. (c) Tachyzoites in a cytospin smear of pleural fluid from a
cat with pneumonia. Giemsa stain. Compare the size of tachyzoites (arrow) with host cells. Giemsa stain. Bar = 10 µm.
(d) Tachyzoites (arrows) and tissue cysts (large arrow) in section of mouse brain. Immunohistochemical stain with
T. ­gondii–specific antibody. Bar = 10 µm.
After the ingestion of tissue cysts by cats, the tissue cyst wall is dissolved by proteolytic enzymes
in the stomach and small intestine. The released bradyzoites penetrate the epithelial cells of the small
intestine and initiate development of numerous generations of asexual and sexual cycles of T. gondii.25
Bradyzoites penetrate the lamina propria of the feline intestine and multiply as tachyzoites. Within a few
hours after infection of cats, T. gondii may disseminate to extraintestinal tissues. T. gondii persists in
intestinal and extraintestinal tissues of cats for at least several months and possibly for the life of the cat.
As the enteroepithelial cycle progresses, T. gondii multiplies profusely in intestinal epithelial cells of
cats (enteroepithelial cycle), and these stages, represented by five distinct morphological types (Types a–e),
are known as schizonts (Figure 12.3d). Several generations of each type are produced, and daughter organisms known as merozoites are formed coincident with the last nuclear division. Merozoites give rise to
gametes, the sexual stages of the organism. The microgamont (male gamont) has two flagella (Figure 12.3e)
and it swims to, enters, and fertilizes the macrogamont (female gamont), forming a zygote. After fertilization, oocyst wall formation begins around the zygote. Oocysts are discharged into the intestinal lumen by
the rupture of intestinal epithelial cells.
Oocysts are environmentally resistant and are formed only in felids, probably in all members of the
Felidae (Figure 12.3g). Cats shed oocysts after ingesting any of the three infectious stages of T. gondii,
that is, tachyzoites, bradyzoites, and sporozoites.25–28 Prepatent periods (time to the shedding of oocysts
after initial infection) and frequency of oocyst shedding vary according to the stage of T. gondii ingested.
Biology of Foodborne Parasites
(f )
FIGURE 12.6 Tissue cysts of T. gondii. Bar = 10 µm. (a) Two tissue cysts (arrows). Note thin cyst wall enclosing bradyzoites. Impression smear of mouse brain. Silver impregnation and Giemsa stain. (b) A tissue cyst freed from mouse brain by
homogenization in saline. Note thin cyst wall (arrow) enclosing many bradyzoites. Unstained. (c) A large tissue cyst in section of rat brain 14 months postinfection. Note thin cyst wall (arrow). H & E stain. (d) A small tissue cyst with intact cyst wall
(arrow) and four bradyzoites (arrowheads) with terminal nuclei adjacent to it. Section of mouse brain 8 months postinfection.
H & E stain. (e) A tissue cyst in section of mouse brain. Note PAS-negative cyst wall (arrow) enclosing many PAS-positive
bradyzoites (arrowheads). The bradyzoites stain bright red with PAS, but they appear black in this photograph. Periodic acidSchiff hematoxylin stain (PASH). (f) An elongated tissue cyst (arrow) in section of skeletal muscle of a mouse. PASH stain.
FIGURE 12.7 TEM of a tissue cyst in brain of a mouse 6 months postinfection. Note thin cyst wall (opposing arrows),
numerous bradyzoites each with a conoid (C), and electron-dense rhoptries (R). Bar = 3.0 µm.
Toxoplasma gondii
Prepatent periods are 3–10 days after ingesting tissue cysts and 19 days or more after ingesting tachyzoites or oocysts.25–27 Less than 50% of cats shed oocysts after ingesting tachyzoites or oocysts, whereas
nearly all cats shed oocysts after ingesting tissue cysts.26 In freshly passed feces, oocysts are unsporulated (noninfective; Figure 12.3f). Unsporulated oocysts are subspherical to spherical and are 10 × 12 µm
in diameter. They sporulate (become infectious) outside the cat within 1–5 days depending on aeration
and temperature. Sporulated oocysts contain two ellipsoidal sporocysts (Figure 12.3g), and each sporocyst contains four sporozoites. The sporozoites are 2 × 6–8 µm in size.
12.3 Biology, Genetics, and Genomics
The nucleus of T. gondii is haploid except during sexual recombination in the gut of the cat. Fourteen
chromosomes are present, encompassing a 65 Mb genome.6,29 Most T. gondii isolates from human and
animal sources in North America, Europe, and Africa have been grouped into one of three clonal lineages, Types I–III,29,30–35 and are biologically and genetically different from isolates from Brazil and
Columbia.36–40 A number of recent studies suggest that only a few ancestral strains have given rise to
the three dominant clonal lineages and the existing genetic diversity seen in various geographic regions
through a process of limited, mostly asexual, recombination.31,34,40,41 Recently, Su et al.40 demonstrated a
biphasic pattern where a few clonal lineages dominate the population in the Northern Hemisphere while,
particularly in South America, populations are much more genetically diverse. Recent genotyping studies of isolates from pigs, lambs, and goats demonstrate that Type II lineage predominates in food animals
in the United States, followed by Type III isolates and atypical genotypes; Type I isolates have rarely
been found in farm animals.35,42–44
12.4 Diagnosis
Diagnosis is made by biologic, serologic, or histologic methods or by combinations of these. Clinical
signs of toxoplasmosis are nonspecific and are not sufficiently characteristic for a definite diagnosis.
Toxoplasmosis in fact mimics several other infectious diseases.
Detection of T. gondii antibody in patients may aid diagnosis. There are numerous serologic procedures available for the detection of humoral antibodies; these include the Sabin–Feldman dye test
(DT), the modified agglutination test (MAT), the indirect hemagglutination test (IHAT), the indirect
fluorescent antibody assay (IFA), the direct agglutination test, the latex agglutination test (LAT), the
enzyme-linked immunosorbent assay (ELISA), and the immunosorbent agglutination assay test (IAAT).
The IFA, IAAT, and ELISA have been modified to detect IgM antibodies.45 The IgM antibodies appear
sooner after infection than the IgG antibodies and disappear faster than IgG antibodies after recovery,
though a small percentage of infected people produce IgG first.45,46 Progress has been made in the diagnosis of human infection with Toxoplasma using PCR.47 Infection has been diagnosed using nested,
stage-specific primers and cerebrospinal fluid from acquired immunodeficiency syndrome (AIDS)
patients with suspected toxoplasmic encephalitis,48,49 in immunocompromised patients undergoing
hematopoietic stem cell transplantation,50 and in suspected cases of fetal toxoplasmosis using amniotic
fluid.51 Improved sensitivity and performance standards for in-house methods and commercially available PCR kits are needed, as recent studies have shown that these PCR tests may not perform well using
experimental or clinical samples.52–54
12.5 Epidemiology
Toxoplasmosis may be acquired by ingestion of oocysts or by ingestion of tissue-inhabiting stages of the
parasite. Contamination of the environment by oocysts is widespread as oocysts are shed by domestic
cats and other members of the Felidae.1,24 Domestic cats are probably the major source of contamination
as oocyst formation is greatest in domestic cats, and cats are extremely common. Widespread natural
Biology of Foodborne Parasites
infection of the environment is possible since a cat may excrete millions of oocysts after ingesting as few
as one bradyzoite or one tissue cyst, and many tissue cysts may be present in one infected mouse.24,55
Sporulated oocysts survive for long periods under most ordinary environmental conditions and even in
harsh environments for months. They can survive in moist soil, for example, for months and even years.1
Oocysts in soil can be mechanically transmitted by invertebrates such as flies, cockroaches, dung
beetles, and earthworms, which can spread oocysts into human food and animal feeds.
Infection rates in cats are determined by the rate of infection in local avian and rodent populations
because cats typically become infected by eating these animals. The more oocysts in the environment,
the more likely it is that prey animals would be infected, and this in turn would increase the infection
rate in cats. In certain areas of Brazil, up to 50% of 6–8-year-old children have antibodies to T. gondii
linked to the ingestion of oocysts from the environment heavily contaminated with T. gondii oocysts.56,57
The largest recorded outbreak of clinical toxoplasmosis in humans in North America was epidemiologically linked to drinking water from a municipal water reservoir in British Columbia, Canada.58,59
This water reservoir was supposedly contaminated with T. gondii oocysts excreted by cougars (Felis
concolor). Although attempts to recover T. gondii oocysts from water samples in the British Columbia
outbreak were unsuccessful, methods to detect oocysts were reported.60 At present, there are no commercial reagents available to reliably detect T. gondii oocysts in the environment.
Widespread infection in aquatic mammals indicates contamination and survival of oocysts in seawater.61,62 Wild populations of southern sea otters have been significantly impacted by exposure to
Toxoplasma oocysts, presumably by eating filter-feeding mollusks in near-shore environments.21,63
Transmission of Toxoplasma from consumption of infected meat products is difficult to quantify,
since meat from infected animals may undergo postharvest treatments such as heating, freezing, salting,
or pumping (injection of water- and salt-based solutions to retard microbial growth) that can render the
parasite nonviable,64,65 and few comprehensive assessments have been completed in meat available for
retail purchase. Complicating matters is the fact that the number of T. gondii organisms in meat from
naturally infected food animals is very low, making the parasite difficult and expensive to detect by
direct methods. It is estimated that as few as one tissue cyst may be present in 100 g of meat.6 In addition,
there is no predilection site for Toxoplasma in meat animals; virtually, all edible portions of an animal
can harbor viable T. gondii tissue cysts,13 and tissue cysts can remain viable in food animals for years.
Beef, chicken, and pork are the main meat types consumed in the United States. In a case-control
study of 148 recently (<6 months) infected individuals, Jones et al.66 identified an elevated risk of infection associated with eating raw ground beef; rare lamb; locally produced cured, dried, or smoked meat;
and raw oysters, clams, or mussels, working with meat, and drinking unpasteurized goat milk.
The relative risk to U.S. consumers of acquiring T. gondii infection from undercooked meat was recently
determined in a nationwide survey of retail chicken, beef, and pork. The survey of 698 retail outlets in
28 metropolitan statistical areas (MSAs as defined by the U.S. Census Bureau) covered 80% of the U.S.
population. Only pork was found to harbor viable T. gondii tissue cysts, which were isolated from 0.38%
of samples (7/2094) by cat bioassay, and 0.57% of pork samples were suspected to be infected based on
positive ELISA results. No beef samples were positive by bioassay or by ELISA, while 1.4% of chickens were positive by ELISA only. The northeastern United States had a higher number of positive pork
samples than other regions of the country, reflecting the higher risk of pig infection due to regional management practices (outdoor vs. confinement rearing17). Thus, while the extent of human infection resulting
from meat sources remains undetermined, the lack of viable organisms in beef and chicken and the low
prevalence of T. gondii infection in market pigs found in this comprehensive study would not account for
the estimated incidence and measured seroprevalence in humans in the United States.
12.6 Pathogenesis and Clinical Features
T. gondii usually parasitizes the host, definitive and intermediate, without producing clinical disease.
Only rarely does it produce severe clinical manifestations. The bradyzoites from the tissue cysts, or sporozoites from the oocyst, penetrate intestinal epithelial cells and multiply in the intestine as tachyzoites
within 24 h of infection. T. gondii may spread first to mesenteric lymph nodes and then to distant organs
Toxoplasma gondii
by invasion of lymphatics and blood and can multiply in virtually any cell in the body. All extracellular
forms of the parasite are directly affected by antibody, but intracellular forms are not. More virulent
strains of Toxoplasma have developed effective defensive mechanisms using ROP18, a rhoptry-associated serine/threonine kinase, to inactivate p47 GTPases, which are generated by the infected cell to
rupture the vacuole containing the parasite, resulting in digestion of the organism.67 It is believed that
cellular factors, including lymphocytes and lymphokines, are more important than humoral factors in
immune-mediated destruction of T. gondii.68–70 Immunity does not eradicate infection. T. gondii tissue
cysts persist for years after acute infection. The fate of tissue cysts is not fully known. Whether bradyzoites can form new tissue cysts directly without transforming into tachyzoites is not known. It has been
proposed that tissue cysts may at times rupture during the life of the host. The released bradyzoites may
be destroyed by the host’s immune responses, or there may be formation of new tissue cysts. In immunosuppressed patients, such as those given large doses of immunosuppressive agents in preparation for
organ transplants and in those with AIDS, rupture of a tissue cyst may result in transformation of bradyzoites into tachyzoites and renewed multiplication. The immunosuppressed host may die from toxoplasmosis unless treated. It is not known how corticosteroids cause relapse, but it is unlikely that they directly
cause rupture of the tissue cysts.
Pathogenicity of T. gondii is determined by the virulence of the strain and the susceptibility of the host
species.71 T. gondii strains may vary in their pathogenicity in a given host. Certain strains of mice are
more susceptible than others, and the severity of infection in individual mice within the same strain may
vary. Mice of any age are susceptible to clinical T. gondii infection.6 However, adult rats do not become
ill, while young rats can die of toxoplasmosis. Adult dogs, like adult rats, are resistant, whereas puppies are fully susceptible to clinical toxoplasmosis. Certain species are genetically resistant to clinical
toxoplasmosis. Cattle and horses are among the hosts more resistant to clinical toxoplasmosis, whereas
certain marsupials and new-world monkeys are highly susceptible to T. gondii infection.1,6 Nothing is
known concerning genetically determined susceptibility to clinical toxoplasmosis in higher mammals,
including humans.
Infection in humans may be congenitally or postnatally acquired. Congenital infection occurs only
when a woman becomes infected during pregnancy. Congenital infections acquired during the first trimester are more severe than those acquired in the second and third trimesters.45,72 While the mother
rarely has symptoms of infection, she does have a temporary parasitemia. Focal lesions develop in the
placenta, and the fetus may become infected. At first, there is generalized infection in the fetus. Later,
infection is cleared from the visceral tissues and may localize in the central nervous system. A wide
spectrum of clinical diseases occur in congenitally infected children.72 Mild disease may consist of
slightly diminished vision, whereas severely diseased children may have the full tetrad of lesions of the
eye, hydrocephalus, convulsions, and intracerebral calcification. Of these, hydrocephalus is the least
common but most significant lesion of toxoplasmosis. So far, the most common sequela of congenital
toxoplasmosis is ocular disease.45,72 The socioeconomic impact of toxoplasmosis in human suffering and
the cost of care of sick children, especially those with mental retardation and blindness, are enormous.73,74
The testing of all pregnant women for T. gondii infection is compulsory in some European countries,
including France and Austria.75,76 The cost benefits of such mass screening are being debated in many
other countries.45,77,78 Recently, Stillwaggon et al.79 provided an extensive guideline for estimating costs
of preventive maternal screening for and the social costs resulting from toxoplasmosis based on studies
in Europe and the United States. While estimating these costs, the value of all resources used or lost
should be considered, including the cost of medical and nonmedical services, wages lost, cost of in-home
care, and indirect costs of psychological impacts borne by the family for lifetime care of a substantially
cognitively impaired child; cost of a fetal death was estimated to be $5 million dollars.
Postnatally acquired infection may be localized or generalized. Infection may occur in any organ.
Oocyst-transmitted infections may be more severe than tissue cyst–induced infections.1,80–84 Enlarged
lymph nodes are the most frequently observed clinical sign of toxoplasmosis in humans (Table 12.1).
Lymphadenopathy may be associated with fever, fatigue, muscle pain, sore throat, and headache.
Although the condition may be benign, its diagnosis is vital in pregnant women because of the risk
to the fetus. In a British Columbia outbreak, of the 100 people who were diagnosed with acute infection, 51 had lymphadenopathy and 20 had retinitis.58,59 Encephalitis is the most important manifestation
Biology of Foodborne Parasites
TABLE 12.1
Frequency of Symptoms in People with Postnatally Acquired Toxoplasmosis
Patients with Symptoms (%)
Stiff neck
Sore throat
Eye pain
Abdominal pain
Atlanta Outbreaka
(35 Patients)
Panama Outbreakb
(35 Patients)
From Teutsch et al. [80].
From Benenson et al. [81].
Not reported.
of toxoplasmosis in immunosuppressed patients as it causes the most severe damage to the patient.1,85
Patients may have headache, disorientation, drowsiness, hemiparesis, reflex changes, and convulsions,
and many become comatose. Encephalitis caused by T. gondii is now recognized with great frequency in
patients treated with immunosuppressive agents.
Toxoplasmosis ranked high on the list of diseases that lead to death of patients with AIDS, and approximately 10% of AIDS patients in the United States and up to 30% in Europe have died from toxoplasmosis.85 In AIDS patients, although any organ may be involved, including the testis, dermis, and the spinal
cord, infection of the brain is most frequently reported. Most AIDS patients suffering from toxoplasmosis have bilateral, severe, and persistent headaches, which respond poorly to analgesics. As the disease
progresses, the headaches may give way to a condition characterized by confusion, lethargy, ataxia, and
coma. The predominant lesion in the brain is necrosis, especially of the thalamus.86 Since the advent of
highly active antiretroviral therapy (HAART) in the mid-1990s, the number of AIDS patients suffering
from toxoplasmic encephalitis has fallen dramatically, at least partially due to the impact of protease
inhibitors used in HAART on Toxoplasma proteases.87–89
12.7 Treatment and Prevention
Sulfadiazine and pyrimethamine (Daraprim) are two drugs widely used in the treatment of toxoplasmosis.90,91 While these drugs have a beneficial action when given in the acute stage of the disease process
when there is active multiplication of the parasite, they will not usually eradicate infection. It is believed
that these drugs have little effect on subclinical infections, but the growth of tissue cysts in mice has been
restrained with sulfonamides. Some other drugs, like diaminodiphenylsulfone, atovaquone, spiramycin,
and clindamycin, are also used to treat toxoplasmosis in difficult cases.
To prevent infection of human beings by T. gondii, the hands of people handling meat should be washed
thoroughly with soap and water before they go to other tasks.1,92 All cutting boards, sink tops, knives, and
other materials coming in contact with uncooked meat should be washed with soap and water. Washing
is effective because the stages of T. gondii in meat are killed by contact with soap and water.1 T. gondii
organisms in meat can be killed by exposure to extreme cold or heat. Tissue cysts in meat are killed by
Toxoplasma gondii
heating the meat throughout to 67°C93 and by cooling to −13°C.94 Toxoplasma in tissue cysts are also
killed by exposure to 0.5 kilorads of gamma irradiation.95 Meat of any animal should be cooked to 67°C
before consumption, and tasting meat while cooking or while seasoning should be avoided.
Pregnant women and immunocompromised patients should avoid contact with cats, soil, and raw
meat. Pet cats should be fed with only dry, canned, or cooked food. The cat litter box should be emptied
every day, preferably not by a pregnant woman. Gloves should be worn while gardening. Vegetables
should be washed thoroughly before eating because they may have been contaminated with cat feces.
Expectant mothers should be aware of the dangers of toxoplasmosis.96,97 At present, there is no vaccine
to prevent toxoplasmosis in humans.
12.8 Future Directions and Trends
Toxoplasmosis continues to be a significant public health problem worldwide. Although toxoplasmosis is
estimated to have a disease burden and economic impact comparable to that of campylobacteriosis and
salmonellosis, there are presently no explicit monitoring programs to screen animals entering the food
chain and no standardized reporting of human toxoplasmosis between different countries. The increasing demand for food safety together with the potential economic impact of legislation aimed at risk
reduction has brought attention to the need for development and standardization of diagnostic tests for
Toxoplasma infection, including assays that allow researchers to assess the role of different transmission
routes in disease epidemiology. Such tests will need to provide an accurate estimate of risks of transmission of Toxoplasma to humans and must perform with comparable specificity and sensitivity across a
range of animal species.
1. Dubey, J.P. and Beattie, C.P., Toxoplasmosis of Animals and Man, CRC Press, Boca Raton, FL, 1988.
2. Dubey, J.P. and Jones, J.L., Toxoplasma gondii infection in humans and animals in the United States, Int.
J. Parasitol., 38, 1257, 2008.
3. Jones, J.L. et al., Toxoplasma gondii infection in the United States: Seroprevalence and risk factors, Am.
J. Epidemiol., 154, 357, 2001.
4. Jones, J.L. et al., Toxoplasma gondii infection in the United States, 1999–2000, Emerg. Infect. Dis., 9,
1371, 2003.
5. Jones, J.L. et al., Toxoplasma gondii infection in the United States, 1999–2004, decline from the prior
decade, Am. J. Trop. Med. Hyg., 77, 405, 2007.
6. Dubey, J.P., Toxoplasmosis of Animals and Humans, 2nd edn., CRC Press, Boca Raton, FL, 2010.
7. Minbaeva, G. et al., Toxoplasma gondii infection in Kyrgyzstan: Seroprevalence, risk factor analysis, and
estimate of congenital and AIDS-related toxoplasmosis, PLoS Negl. Trop. Dis., 7(2), e2043, 2013.
8. Jones, J.L. and Holland, G.N., Annual burden of ocular toxoplasmosis in the United States, Am. J. Trop.
Med. Hyg., 82, 464, 2010.
9. Batz, M.B., Hoffmann, S., and Morris Jr. J.G., Ranking the disease burden of 14 pathogens in food
sources in the United States using attribution data from outbreak investigations and expert elicitation,
J. Food Prot., 75(7), 1278–1291, 2012.
10. Hoffmann, S., Batz, M.B., and Morris, Jr. J.G., Annual cost of illness and quality-adjusted life year losses
in the United States due to 14 foodborne pathogens, J. Food Prot., 75(7), 1292, 2012.
11. Pedersen, M.G. et al., Toxoplasma gondii infection and self-directed violence in mothers, Arch. Gen.
Psychiatry, 69(11), 1123, 2012.
12. Torrey, E.F., Bartko, J.J., and Yolken, R.H., Toxoplasma gondii and other risk factors for schizophrenia:
An update, Schizophr. Bull., 38(3), 642, 2012.
13. Dubey, J.P. et al., Distribution of Toxoplasma gondii tissue cysts in commercial cuts of pork, J. Am. Vet.
Med. Assoc., 188, 1035, 1986.
14. Mead, P.S. et al., Food-related illness and death in the United States, Emerg. Infect. Dis., 5, 607, 1999.
15. Roghmann, M.C. et al., Decreased seroprevalence for Toxoplasma gondii in seventh day adventists in
Maryland, Am. J. Trop. Med. Hyg., 60, 790, 1999.
Biology of Foodborne Parasites
16. Scallan, E. et al., Foodborne illness acquired in the United States-major pathogens, Emerg. Infect. Dis.,
17, 7, 2011.
17. Dubey, J.P. et al., Prevalence of viable Toxoplasma gondii in beef, chicken and pork from retail meat
stores in the United States: Risk assessment to consumers, J. Parasitol., 91, 1082, 2005.
18. Hill, D. et al., Identification of a sporozoite-specific antigen from Toxoplasma gondii, J. Parasitol., 97,
328, 2011.
19. Boyer, K. et al., Unrecognized ingestion of Toxoplasma gondii oocysts leads to congenital toxoplasmosis
and causes epidemics in North America, Clin. Infect. Dis., 53(11), 1081, 2011.
20. Levy, J.K. and Crawford, P.C., Humane strategies for controlling feral cat populations, J. Am. Vet. Med.
Assoc., 225, 1354, 2004.
21. Miller, M.A. et al., Coastal freshwater runoff is a risk factor for Toxoplasma gondii infection of southern
sea otters (Enhydra lutris nereis), Int. J. Parasitol., 32(8), 997, 2002.
22. Conrad, P.A. et al., Transmission of Toxoplasma: Clues from the study of sea otters as sentinels of
Toxoplasma gondii flow into the marine environment, Int. J. Parasitol., 35(11–12), 1155, 2005.
23. Hill, D.E., Chirukandoth, S., and Dubey, J.P., Biology and epidemiology of Toxoplasma gondii in man
and animals, Anim. Health Res. Rev., 6(1), 41, 2005.
24. Frenkel, J.K., Dubey, J.P., and Miller, N.L., Toxoplasma gondii in cats: Fecal stages identified as coccidian oocysts, Science, 167(3919), 893, 1970.
25. Dubey, J.P. and Frenkel, J.K., Cyst-induced toxoplasmosis in cats, J. Protozool., 19(1), 155, 1972.
26. Dubey, J.P. and Frenkel, J.K., Feline toxoplasmosis from acutely infected mice and the development of
Toxoplasma cysts, J. Protozool., 23(4), 537, 1976.
27. Dubey, J.P., Infectivity and pathogenicity of Toxoplasma gondii oocysts for cats, J. Parasitol., 82(6), 957,
28. Dubey, J.P., Tachyzoite-induced life cycle of Toxoplasma gondii in cats, J. Parasitol., 88(4), 713, 2002.
29. Khan, A. et al., Recent transcontinental sweep of Toxoplasma gondii driven by a single monomorphic
chromosome, Proc. Natl. Acad. Sci. USA, 104(37), 14872, 2007.
30. Dardé, M.L., Bouteille, B., and Perstreal, M., Isoenzyme analysis of 35 Toxoplasma gondii isolates and
the biological and epidemiologic implications, J. Parasitol., 78, 909, 1992.
31. Howe, D.K. and Sibley, L.D., Toxoplasma gondii comprises three clonal lineages: Correlation of parasite
genotype with human disease, J. Infect. Dis., 172, 1561, 1995.
32. Ajzenberg, D. et al., Microsatellite analysis of Toxoplasma gondii shows considerable polymorphism
structured into two main clonal groups, Int. J. Parasitol., 32, 27, 2002.
33. Ajzenberg, D. et al., Genotype of 86 Toxoplasma gondii isolates associated with human congenital toxoplasmosis, and correlation with clinical findings, J. Infect. Dis., 186, 684, 2002.
34. Su, C. et al., Recent expansion of Toxoplasma through enhanced oral transmission, Science, 299(5605),
414, 2003.
35. Velmurugan, G.V., Su, C., and Dubey, J.P., Isolate designation and characterization of Toxoplasma gondii
isolates from pigs in the United States, J. Parasitol., 95, 95, 2009.
36. Dubey, J.P. et al., Biological and genetic characterization of Toxoplasma gondii isolates from chickens
(Gallus domesticus) from São Paulo, Brazil: Unexpected findings, Int. J. Parasitol., 32, 99, 2002.
37. Dubey, J.P. et al., Prevalence of Toxoplasma gondii in dogs from Colombia, South America and genetic
characterization of T. gondii isolates, Vet. Parasitol., 145, 45, 2007.
38. Lehmann, T. et al., Globalization and the population structure of Toxoplasma gondii, Proc. Natl. Acad.
Sci. USA, 103, 11423, 2006.
39. Dubey, J.P. and Su, C., Population biology of Toxoplasma gondii: What’s out and here did they come
from, Mem. Inst. Oswaldo Cruz, 104, 190, 2009.
40. Su, C. et al., Globally diverse Toxoplasma gondii isolates comprise six major clades originating from a
small number of distinct ancestral lineages, Proc. Natl. Acad. Sci. USA, 109(15), 5844, 2012.
41. Grigg, M.E. et al., Success and virulence in Toxoplasma as the result of sexual recombination between
two distinct ancestries, Science, 294(5540), 161, 2001.
42. Dubey, J.P. et al., Endemic toxoplasmosis in pigs on a farm in Maryland: Isolation and genetic characterization of Toxoplasma gondii, J. Parasitol., 94, 36, 2008.
43. Dubey, J.P. et al., High prevalence and abundant atypical genotypes of Toxoplasma gondii isolated from
lambs destined for human consumption in the USA, Int. J. Parasitol., 38, 999, 2008.
Toxoplasma gondii
44. Dubey, J.P. et al., High prevalence and genotypes of Toxoplasma gondii isolated from goats, from a retail
meat store, destined for human consumption in the USA, Int. J. Parasitol., 41, 827, 2011.
45. Remington, J.S., McLeod, R., and Desmonts, G., Toxoplasmosis. In: Remington, J.S., and Klein, J., Eds.
Infectious Diseases of the Fetus and Newborn Infant. W.B. Saunders, Philadelphia, PA, p. 140, 1995.
46. Fricker-Hidalgo, H. et al., Toxoplasma seroconversion with negative or transient immunoglobulin M in
pregnant women: Myth or reality? A French multicentre retrospective study, J. Clin. Microbiol., E-pub
April 24, 2013. PMID: 23616461.
47. Rahumatullah, A., Khoo, B.Y., and Noordin, R., Triplex PCR using new primers for the detection of
Toxoplasma gondii, Exp. Parasitol., 131(2), 231, 2012.
48. Contini, C. et al., The role of stage-specific oligonucleotide primers in providing effective laboratory
support for the molecular diagnosis of reactivated Toxoplasma gondii encephalitis in patients with AIDS,
J. Med. Microbiol., 51(10), 879, 2002.
49. Joseph, P. et al., Optimization and evaluation of a PCR assay for detecting toxoplasmic encephalitis in
patients with AIDS, J. Clin. Microbiol., 40(12), 4499, 2002.
50. Lewis, J.S. Jr et al., PCR for the diagnosis of toxoplasmosis after hematopoietic stem cell transplantation,
Expert Rev. Mol. Diagn., 2(6), 616, 2002.
51. Antsaklis, A. et al., Prenatal diagnosis of congenital toxoplasmosis, Prenat. Diagn., 22(12),1107, 2002.
52. Hill, D.E. et al., Comparison of detection methods for Toxoplasma gondii in naturally and experimentally
infected swine, Vet. Parasitol., 141(1–2), 9, 2006.
53. Morelle, C. et al., Comparative assessment of a commercial kit and two laboratory-developed PCR
assays for molecular diagnosis of congenital toxoplasmosis, J. Clin. Microbiol., 50(12), 3977, 2012.
54. Mikita, K. et al., The utility of cerebrospinal fluid for the molecular diagnosis of toxoplasmic encephalitis, Diagn. Microbiol. Infect. Dis., 75(2), 155, 2013.
55. Dubey, J.P., Oocyst shedding by cats fed isolated bradyzoites and comparison of infectivity of bradyzoites of the VEG strain Toxoplasma gondii to cats and mice, J. Parasitol., 87(1), 215, 2001.
56. Bahia-Oliveira, L.M. et al., Highly endemic, waterborne toxoplasmosis in north Rio de Janeiro state,
Brazil, Emerg. Infect. Dis., 9(1), 55, 2003.
57. Dubey, J.P. et al., Toxoplasmosis in humans and animals in Brazil: High prevalence, high burden of disease, and epidemiology, Parasitology, 139(11), 1375, 2012.
58. Aramini, J.J., Stephen, C., and Dubey, J.P., Toxoplasma gondii in Vancouver Island cougars (Felis concolor vancouverensis): Serology and oocyst shedding, J. Parasitol., 84, 438, 1998.
59. Aramini, J.J. et al., Potential contamination of drinking water with Toxoplasma gondii oocysts, Epidem.
Infec., 122, 305, 1999.
60. Isaac-Renton, J. et al., Detection of Toxoplasma gondii oocysts in drinking water, App. Envir. Microbio.,
64, 2278, 1998.
61. Cole, R.A. et al., Biological and molecular characterizations of Toxoplasma gondii strains obtained from
southern sea otters (Enhydra lutris nereis), J. Parasitol., 86, 526, 2000.
62. Lindsay, D.S. and Dubey, J.P., Long-term survival of Toxoplasma gondii sporulated oocysts in seawater,
J. Parasitol., 95(4), 1019, 2009.
63. Miller, M.A. et al., Type X Toxoplasma gondii in a wild mussel and terrestrial carnivores from coastal
California: New linkages between terrestrial mammals, runoff and toxoplasmosis of sea otters, Int.
J. Parasitol., 38(11), 1319, 2008.
64. Hill, D.E. et al., Effect of commonly used enhancement solutions on the viability of Toxoplasma gondii
tissue cysts in pork loin, J. Food Protect., 67, 2230, 2004.
65. Hill, D.E. et al., Effect of time and temperature on the viability of Toxoplasma gondii tissue cysts in
enhanced pork loin, J. Food Protect., 69, 1961, 2006.
66. Jones, J.L. et al., Risk factors for Toxoplasma gondii infection in the United States, Clin. Inf. Dis., 49,
878, 2009.
67. Fentress, S.J. and Sibley, L.D., The secreted kinase ROP18 defends Toxoplasma’s border, Bioessays,
33(9), 693, 2011.
68. Gigley, J.P., Fox, B.A., and Bzik, D.J., Cell-mediated immunity to Toxoplasma gondii develops primarily by
local Th1 host immune responses in the absence of parasite replication, J. Immunol., 182(2), 1069, 2009.
69. Vouldoukis, I. et al., IgE mediates killing of intracellular Toxoplasma gondii by human macrophages
through CD23-dependent, interleukin-10 sensitive pathway, PLoS One, 6(4), e18289, 2011.
Biology of Foodborne Parasites
70. Koshy, A.A. et al., Toxoplasma co-opts host cells it does not invade, PLoS Pathog., 8(7), e1002825, 2012.
71. Hunter, C.A. and Sibley, L.D., Modulation of innate immunity by Toxoplasma gondii virulence effectors,
Nat. Rev. Microbiol., 10(11), 766, 2012.
72. Desmonts, G. and Couvreur, J., Congenital toxoplasmosis. A prospective study of 378 pregnancies,
NEJM, 290, 1110, 1974.
73. Roberts, T. and Frenkel, J.K., Estimating income losses and other preventable costs caused by congenital
toxoplasmosis in people in the United States, JAVMA, 196, 249, 1990.
74. Roberts, T., Murrell, K.D., and Marks, S., Economic losses caused by foodborne parasitic diseases,
Parasitol. Today, 10, 419, 1994.
75. Thiebaut, R. et al., Effectiveness of prenatal treatment for congenital toxoplasmosis: A meta-analysis of
individual patients’ data, Lancet, 369(9556), 115, 2007.
76. Petersen, E., Prevention and treatment of congenital toxoplasmosis, Expert Rev. Anti. Infect. Ther., 5(2),
285, 2007.
77. Cortina-Borja, M. et al., Prenatal treatment for serious neurological sequelae of congenital toxoplasmosis: An observational prospective cohort study, PloS Med., 7(10), pii:e1000351, 2010.
78. Remington, J.S. et al., Toxoplasmosis. In: Remington, J.S. and Klein, J. (Eds), Infectious Diseases of the
Fetus and Newborn Infant. W.B. Saunders, Philadelphia, PA, p. 205, 2001.
79. Stillwaggon, E. et al., Maternal serologic screening to prevent congenital toxoplasmosis: A decisionanalytic economic model, PLoS Negl. Trop. Dis., 5, e1333, 2011.
80. Teutsch, S.M. et al., Epidemic toxoplasmosis associated with infected cats, NEJM, 300, 695, 1979.
81. Benenson, M.W. et al., Oocyst-transmitted toxoplasmosis associated with ingestion of contaminated
water, NEJM, 307, 666, 1982.
82. Smith J.L., Documented outbreaks of toxoplasmosis: Transmission of Toxoplasma gondii to humans,
J. Food Protect., 56, 630, 1993.
83. Bowie, W.R. et al., Outbreak of toxoplasmosis associated with municipal drinking water, Lancet, 350,
173, 1997.
84. Burnett, A.J. et al., Multiple cases of acquired toxoplasmosis retinitis presenting in an outbreak,
Ophthalmology, 105, 1032, 1998.
85. Luft, B.J. and Remington, J.S., Toxoplasmic encephalitis in AIDS, Clin. Infec. Dis., 15, 211, 1992.
86. Renold, C. et al., Toxoplasma encephalitis in patients with the acquired immunodeficiency syndrome,
Medicine, 71, 224, 1992.
87. Palella, F.J., Declining morbidity and mortality among patients with advanced human immunodeficiency
virus infection, N. Engl. J. Med., 13, 853, 1998.
88. Pozio, E., Highly active antiretroviral therapy and opportunistic protozoan infections, Parasitologia,
46(1–2), 89, 2004.
89. Pozio, E. and Morales, M.A., The impact of HIV-protease inhibitors on opportunistic parasites, Trends
Parasitol., 21(2), 58, 2005.
90. Guerina, N.G. et al., Neonatal serologic screening and early treatment for congenital Toxoplasma gondii
infection. The New England Regional Toxoplasma Working Group, NEJM, 330, 1858, 1994.
91. Chirgwin, K. et al., Randomized phase II trial of atovaquone with pyrimethamine or sulfadiazine for
treatment of toxoplasmic encephalitis in patients with acquired immunodeficiency syndrome: ACTG
237/ANRS 039 study, Clin. Infec. Dis., 34, 1243, 2002.
92. Lopez, A. et al., Preventing congenital toxoplasmosis, MMWR, 49, 59, 2000.
93. Dubey, J.P. et al., Effect of high temperature on infectivity of Toxoplasma gondii tissue cysts in pork,
J. Parasitol., 76(2), 201, 1990.
94. Kotula, A.W. et al., Effect of freezing on infectivity of Toxoplasma gondii tissue cysts in pork, J. Food
Prot., 54, 687, 1991.
95. Dubey, J.P. and Thayer, D.W., Killing of different strains of Toxoplasma gondii tissue cysts by irradiation
under defined conditions, J. Parasitol., 80, 764, 1994.
96. Foulon, W., Naessens, A., and Derde, M.P., Evaluation of the possibilities for preventing congenital toxoplasmosis, Am. J. Perinatol., 11, 57, 1994.
97. Foulon, W., Naessens, A., and Ho-Yen, D., Prevention of congenital toxoplasmosis, J. Perinatal Med., 28,
337, 2000.
Trypanosoma cruzi
Karen Signori Pereira, Flávio Luís Schmidt, Rodrigo Labello Barbosa,
and Luiz Augusto Corrêa Passos
13.1 Introduction................................................................................................................................... 223
13.2 Morphology and Classification..................................................................................................... 224
13.3 Biology, Genetics, and Genomics................................................................................................. 225
13.4 Diagnosis and Typing.................................................................................................................... 226
13.5 Epidemiology and Molecular Epidemiology................................................................................ 227
13.6 Pathogenesis and Clinical Features.............................................................................................. 229
13.7 Treatment and Prevention............................................................................................................. 230
References............................................................................................................................................... 230
13.1 Introduction
American trypanosomiasis (Chagas disease) was discovered in 1909 by Brazilian researcher Carlos
Ribeiro Justiniano das Chagas (1878–1934) in a municipality of the state of Minas Gerais called Lassance.
On April 14, 1909, while examining a feverish 2-year-old child named Berenice, Carlos Chagas discovered in her blood the same protozoon found in “barbers” (hematophagous hemipteran insects) and
several species of animals. Berenice is considered the first described clinical case of Chagas disease in
humans.1,2 The disease involves a complex biological cycle (protozoan life cycle relies on two types of
hosts) and transmission of Trypanosoma cruzi.
The protozoan genus name is derived from the Greek trypano (= drill, auger) and soma (= body)
because of their corkscrew-like motion. The specific epithet was an honor to Oswaldo Cruz, a renowned
microbiologist and Carlos Chagas’ mentor.3
Chagas disease is among the most important parasitic diseases. The World Health Organization
(WHO) estimates that about seven to eight million people worldwide are infected with T. cruzi, mostly
in Latin America.4,5 In South America, especially Brazil, Ecuador, Chile, Paraguay, Uruguay, Bolivia,
Argentina, and Venezuela, Chagas disease is a severe and alarming public health problem. Since it
affects mainly people in developing countries, it is a neglected disease, receiving little funding for diagnosis, treatment, and prevention. However, in the past decades, it has been increasingly detected in the
United States, Canada, and many European and some Western Pacific countries.5
More than a century after the disease was discovered and described, a cure is still unavailable, and
many aspects of the pathogen and disease still need elucidation.
Historically, man became infected mainly in the rural and periurban areas of Latin America, when
the natural environment was changed or destroyed, as vectorial transmission was the most important
route of T. cruzi transmission. Nowadays, oral route transmission, notably associated with ingestion of
contaminated food, has shown increasing importance in endemic countries.5–7
Foodborne transmission of Chagas disease is mainly associated with the ingestion of contaminated
breast milk, raw milk, water, or fruit and leaf vegetable juices.8,9 Often, insect vectors are crushed
during food preparation, or the food is contaminated by vector or reservoir (opossum’s) excrements.10
Biology of Foodborne Parasites
Additionally, foodborne transmission can stem from handling or ingesting raw, undercooked, or smoked
meat from infected mammals,11 ingesting armadillo and opossum blood used as medication by Colombian
Amazon medicine, or engaging in primitive habits, like eating triatomines.12
13.2 Morphology and Classification
T. cruzi is a hemoflagellate that may assume distinct morphologic forms. Among cell characteristics,
underlying the plasma membrane is a system of microtubules, the subpellicular microtubular network,
which forms a spiral framework just beneath the surface of the pellicle and provides limited structural
support. A flagellum arises from a basal body in close association with a prominent kinetoplast, forming a kinetoplast–basal body complex. Kinetoplast is rich in DNA that resembles mitochondrial DNA
of other organisms, being composed of a limited number of nucleotides arranged in linked circlets.
Kinetoplast DNA (kDNA) appears to be responsible for the elaboration of mitochondria in many of the
forms as well as for the metamorphosis of one stage to another.2,13
During the life cycle and involving different hosts (vertebrates and invertebrates), T. cruzi may assume
different morphologic forms, according to the position of the kinetoplast and basal body to the nucleus
and the emergence of the flagellum.13,14 Four morphologic forms are represented in Figure 13.1: (a) amastigote, which is ovoid, characterized by a single prominent nucleus, a very short flagellum, and usually
develops in vertebrate host cells; (b) promastigote, which is more elongated than amastigote and the long
flagellum is free anteriorly and serves the function of both locomotion through the medium and attachment to the insect gut wall and occurs only in the insect vector; (c) epimastigote, which is an elongated
cell and the basal body complex is situated more posteriorly but remains anterior to the nucleus; and (d)
trypomastigote, which can have two types (the long, slender trypomastigote and stumpy trypomastigote),
both are characterized by short or no free flagellum, elongation of the undulating membrane and flagellum, and migration of the basal body complex to a site posterior to the nucleus.13
T. cruzi is a protozoan (Protista kingdom and Protozoa subkingdom) of the Sarcomastigophora phylum, Mastigophora subphylum, Kinetoplastida order, Trypanosomatidae family, and Trypanosoma
genus.3 According to Hamilton and Stevens,15 the taxonomy of the genus Trypanosoma is based on
comparisons of morphology, life cycle, and disease data. The most commonly used taxonomy, proposed
by Hoare in 1972, divides mammalian Trypanosoma spp. into two sections: Stercoraria and Salivaria.
T. cruzi is classified in section Stercoraria (transmission is through contact with infective forms in the
feces, generally from insect vector), subgenus Schyzotrypanum.
The family Trypanosomatidae also includes other human pathogens, Trypanosoma brucei (section
Salivaria, subgenus Trypanozoon), the etiological agent of African sleeping sickness, and Leishmania
species, which cause cutaneous and visceral leishmaniasis.16
Basal body
FIGURE 13.1 Schematic forms of T. cruzi: (a) amastigote, (b) promastigote, (c) epimastigote, (d) trypomastigote.
Trypanosoma cruzi
13.3 Biology, Genetics, and Genomics
T. cruzi life cycle is complex, involving invertebrates and vertebrate hosts, and is classified as the heteroxenic type: a phase of intracellular multiplication in vertebrate host (mammals) and an extracellular
multiplication in gut insect vector.2
The invertebrate host (intermediary host) is a hematophagous hemipteran insect whose common name
in Brazil is “the barber.” The vertebrate hosts (definitive host) are mammals such as marsupials and
rodents.17 Some researchers18,19 have demonstrated the importance of common opossum (Didelphis marsupialis) as a T. cruzi reservoir. These authors found that mice became infected after ingesting foods
contaminated with the marsupial’s excrements.
In insects (intermediate host), T. cruzi multiplies in the digestive tract, reaching its infectious stage,
the metacyclic trypomastigote, in the terminal portion of the intestine, where they are excreted with the
insect feces. In mammals, the parasite circulates in the blood and lodges mainly in muscle tissue, such
as the heart, where it multiplies. The parasitic forms of the protozoan reproduce asexually by binary
The protozoan is transmitted from one vertebrate host to another when metacyclic trypomastigote
forms are inoculated to the definitive host or when infected fecal material (from infected insect) is
rubbed into the bite wound, eyes, or mucous membranes.13
The insects become infected by sucking blood from infected mammals (including humans) that have
circulating parasites (trypomastigote forms). In the digestive tract of insect vectors, the trypomastigotes
differentiate into epimastigotes (multiplicative form) and then to metacyclic trypomastigotes in the final
portion of the intestine. Infection of mammals occurs when they come into contact with the infective
metacyclic forms of the parasite that are eliminated with the feces of triatomines after feeding.3 A simplified T. cruzi life cycle is shown in Figure 13.2.
Triatomine insects
Insects bite mammals
ingesting contaminated
blood containing the
Human, opossum,
armadillo, etc.
circulating in the blood
without multiplication
multiply by binary
fission in insect midgut
Cellular internallization
(heart, esophagus, gut cells)
and multiplication (asexual
reproduction) as amastigotes
in insect hindgut
Amastigotes present
in the tissue cells
present in insect
and urine
Insects bite
parasite in feces
Cells die and protozoa are
released into the
FIGURE 13.2 Schematic classic T. cruzi life cycle including events that occur in different hosts.
Biology of Foodborne Parasites
The vectors of Chagas disease are insects of the order Hemiptera, family Reduviidae, and subfamily
Triatominae, whose anterior coriaceous wings give them high dispersal ability. Hemipteran insects, or
true bugs, are classified according to their eating habits; some are phytophagous, some are predators, and
some are hematophagous. Different species can be easily recognized by the shape of their mouth parts.
All hematophagous bugs of the genera Panstrongylus, Rhodnius, and Triatoma are potential vectors
of T. cruzi. The main vector in all Latin America is Triatoma infestans, but other species are also epidemiologically important, including Panstrongylus megistus, Rhodnius pictipes, Triatoma brasiliensis,
Triatoma pseudomaculata, and Triatoma sordida. About 140 triatomine species are described and recognized as valid taxa, with Brazil having the greatest diversity with 61 species.20,21
In addition to vector-borne transmission, the protozoan can also be transmitted by blood transfusion,
transplant of infected organs, laboratory accidents, intercourse, congenital transmission (vertical), and
oral transmission. Another transmission possibility can occur when hunters with wounded hands can
become infected when dealing with infected animals. Oral route transmission can occur in several situations: animals ingesting infected bugs, cannibalism among different species of animals, and ingestion
of food contaminated with feces or urine of infected bugs.2 Foodborne transmission has become an
important route in recent years, especially in Brazilian Amazon region.
13.4 Diagnosis and Typing
The parasite can be detected during the acute phase with a microscope. The trypomastigote forms
can be seen in whole blood, thin blood smear, buffy coat, or thick blood smear stained with Giemsa.
Microhematocrit (Strout’s method) is the reference method. Amastigotes may be found in biopsy specimens stained with hematoxylin and eosin (H&E) or Giemsa.22,23
In the chronic phase, diagnosis relies mainly on xenodiagnoses (in which uninfected triatomine bugs
are fed with patient blood and their gut content examined for parasites 4 weeks later), blood cultures,
inoculation of mice, and cultures in appropriate media such as Novy-MacNeal-Nicolle (NNN) medium
or liver infusion tryptose (LIT) medium.2,22,23
Serological techniques commonly used include enzyme-linked immunosorbent assay (ELISA), indirect hemagglutination, and indirect immunofluorescence tests. However, serologic testing is not very
useful in diagnosing Chagas disease during the acute phase of infection. Although the detection of anti–
T. cruzi immunoglobulin M (IgM) could be used, these assays are not widely available and have not been
standardized.3 In the chronic phase, diagnosis is mainly based on serological testing, but the test results
are sometimes difficult to interpret. The WHO defines a case of Chagas disease as any individual with at
least two positive results out of three serological exams using different techniques.3,4
The use of rapid tests is important for large-scale screening, but their sensitivity is currently too low
for widespread adoption as a first-line diagnostic tool.23
Posttreatment follow-up is currently based on decline or loss of specific IgG antibodies against T. cruzi.
Definitive posttherapeutic reversion and a long period of low serological titers are considered criteria of
cure. This conversion is different among children, adults, and disease phases.23,24
The advent of molecular biology introduces many tools that are more sensitive, accurate, and sometimes rapid. Molecular methods have higher sensitivity than other parasitological methods and, therefore, could be useful to confirm diagnosis in cases of inconclusive serology and as an auxiliary method
to monitor treatment. It is important to point out that molecular methods can identify treatment failure
based on positive detection of T. cruzi DNA, but not treatment success, because even repeated negative
results do not necessarily indicate parasitological cure. At best, such results indicate the absence of
parasite DNA at the time of the test. However, PCR-based methods are not helpful in routine diagnosis
because of poor standardization, potential cross contamination, variable results across laboratories and
countries, and the need for special laboratory facilities.3,25,26
Nested PCR assays provide higher sensitivity than single-run PCR assays and have already been used
in supplementary diagnosis of Chagas disease. However, some disadvantages are time consuming and
false-positive results due to contamination.27,28 In contrast, real-time PCR technology uses fluorescent
labels for continuous monitoring of amplification throughout the reaction. The main advantages are
Trypanosoma cruzi
reduced risk of amplicon carry-over contamination, and shorter turnaround times (amplification and
detection in one step), and potential quantitation.28,29
These molecular biological tools can be used efficiently for many purposes, for example, quality control of blood, organs, and tissues for transplantation, controlling food quality to prevent sales of contaminated foods, investigation of outbreaks, and conducting case studies or ecoepidemiology of triatomines
and infected reservoirs.
On the other hand, according to Zingales,30 the biological heterogeneity of T. cruzi isolates (morphology, DNA content, virulence, pathogenicity, drug susceptibility, and other parameters) is amply documented in the literature, and since the 1970s, these observations have stimulated the search for molecular
markers that could correlate the genotype of the parasite with phenotypic diversity. Therefore, T. cruzi is
now a favorite model for studies of molecular epidemiology and population genetics.
Initial studies of T. cruzi population genetics were based on analysis of isoenzyme electrophoretic
­profiles, historically used to explore the genetic diversity of microorganisms. Such studies revealed the
presence of different group-denominated zymodemes.30 Early studies of Miles et al.31,32 showed the presence of three different T. cruzi zymodemes. Later, Tibayrenc and Ayala analyzed a larger number of
genetic loci in a larger number of T. cruzi isolates and increased diversity to 43 zymodemes.35
According to Macedo et al.,33 the first DNA polymorphism studies in T. cruzi were published in 1980
by Morel et al. who reported restriction fragment length polymorphism (RFLP) typing of minicircles
of kDNA. Parasite populations displaying identical or very similar kDNA minicircle restriction patterns were denominated schizodemes. The minicircle RFLP analysis revealed an unexpected amount of
genetic diversity for T. cruzi, demonstrating that linkage disequilibrium extended even between nuclear
and mitochondrial genome compartments.33–35
Despite genetic variation described for the protozoan, T. cruzi was divided into two major phylogenetic
lineages: T. cruzi I and T. cruzi II. Standardized PCR assays targeted the D7 divergent domain of the
large subunit (LSU) rDNA and mini-exon intergenic region, allowing rapid molecular typing, which was
widely used by the community for epidemiological investigation. In these assays, two sizes of amplicons
are each observed in LSU (110 and 125 bp) and mini-exon (350 and 300 bp) products.30,36 Currently,
T. cruzi is divided into six groups (T. cruzi I–IV), each denominated discrete typing unit (DTU). DTU
is defined by a set of isolates that are genetically similar and can be identified by molecular and immunologic markers.30
El-Sayed et al.37 sequenced the whole T. cruzi genome: a diploid genome containing a predicted
22,570 genes, of which 12,570 represent allelic pairs. Over 50% of the genome consists of repeated
13.5 Epidemiology and Molecular Epidemiology
American trypanosomiasis or Chagas disease is widely distributed in Central and South Americas,
extending from the south of the United States to Argentina.5 Although its prevalence in the Americas
has decreased roughly 70% around the year 2000,38 a 2013 WHO estimate indicates that about seven to
eight million people worldwide are infected with T. cruzi.5
Chagas disease originated millions of years ago as an enzootic infection of wild animals. Since
humans invaded wild ecotopes, Chagas disease became an anthropozoonosis. While evidence
of human infection has been found in mummies up to 9000 years old, endemic Chagas disease
became established as a zoonosis only in the last 200–300 years, as triatomines adapted to domestic
In nature, T. cruzi maintains wild, peridomestic, and domestic cycles. Domestic cycle is maintained
by means of domesticated triatomines that transmit the infection from domestic animals (e.g., dogs, cats,
domestic rats, and mice) to humans and between humans as well. The wild cycle is enzootic and is maintained by triatomines and wild animals (e.g., opossums and armadillos). Peridomestic cycle originates
from the wild cycle and maintains the infection among domestic animals in areas surrounding human
dwellings through the action of peridomestic triatomines. It is important to highlight exchanges between
different cycles; dogs and cats can hunt wild animals, and wild animals, such as rats and opossums,
Biology of Foodborne Parasites
T. cruzi
FIGURE 13.3 Exchanges between different T. cruzi ecological cycles. (Adapted from Coura, J.R. and Dias, J.C.P., Mem.
Inst. Oswaldo Cruz, 104(Suppl. I), 31, 2009.)
sometimes invade areas surrounding human dwellings (Figure 13.3). Opossums (D. marsupialis)
are the most important wild T. cruzi reservoir.17,39 Among triatomines in Brazil, five have particular
­epidemiological importance because they are domesticated: T. infestans (a strictly domestic species),
Panstrongylus megistus, T. brasiliensis, T. pseudomaculata, and T. sordida; other species are wild and
maintain a natural cycle only in wild mammals.4,11,39
In recent years, there has been a “globalization” of Chagas disease due to legal and illegal immigrations from the endemic countries of Latin America to nonendemic countries in North America,
Europe, Asia, and Oceania, in particular the United States, Canada, Spain, France, Switzerland,
Japan, emerging Asian countries, and Australia. These migrations have created new epidemiological
and public health problems for the countries that have received infected immigrants. These problems include risks of transfusion and congenital transmission, as well as a need for medical care for
Chagas patients and additional quality controls in blood banks in countries with little experience in
this subject.39,40
On the other hand, despite the success of programs created for reducing vector-borne transmissions
and the fewer recorded mother-to-child, transfusion-associated, sexual, laboratory-associated, or organ
transplant–associated transmissions, the disease will continue to exist in Brazil in the next decades
because of the great number of chronic Chagas elderly and because of the recent changes in epidemiology, namely, the growing importance of foodborne transmission through ingestion of the metacyclic
trypomastigote forms, given their ability to survive the gastric juice. Foods may be contaminated with
metacyclic trypomastigote of T. cruzi by feces from infected triatomines or with the infected bugs themselves that are inadvertently ground into food during preparation, ingestion of undercooked meat or
blood from an infected animal, and contamination of food or drink with fluids (urine, blood, etc.) from
an infected marsupial.7,8,23,39–43
Soon after the discovery of trypanosomiasis, Carlos Chagas and Oswaldo Cruz described the first case
of foodborne transmission of the disease when black-tufted marmosets (Callithrix penicillata) acquired
the parasite by predation, after being placed in a cage with infected insects. At the time, the hypothesis
was that the black-tufted marmosets became infected by ingesting the insects.12 Figure 13.4 summarizes
some major elements associated with contamination of food by T. cruzi.
Regarding T. cruzi molecular epidemiology, several acute cases of human Chagas disease (commonly
foodborne outbreaks) have been reported in the Amazon region, most of them by T. cruzi I. Moreover,
molecular epidemiological studies indicate that T. cruzi I predominates in wild reservoirs, whereas
T. cruzi II is responsible for Chagas disease in domestic cycle; connection between the two cycles is
mostly through insect vectors that carry T. cruzi II.30,39,44
Trypanosoma cruzi
Didelphis marsupialis
Triatomines, “barbers,”
“kissing bugs”
metacyclic trypomastigote T. cruzi
FIGURE 13.4 Major elements associated with contamination of food by T. cruzi.
13.6 Pathogenesis and Clinical Features
The pathogenesis of Chagas disease is largely due to the immune reactions to the parasite and may present as acute, chronic, or asymptomatic. The disease has two distinct phases: the initial or acute phase,
often asymptomatic or oligosymptomatic, characterized by the presence of the trypomastigote in host
blood, and the second or chronic phase, characterized by lesions in the heart and/or digestive tissues of
the host and the difficulty in detecting circulating protozoans.45,46
There may also be an asymptomatic or latency phase, where there are no obvious clinical symptoms.
Some Chagas patients remain in this phase without ever reaching the chronic phase of the disease.45,46
Individuals in the chronic phase may develop heart failure, megaesophagus, and megacolon.47
Chagas disease is the most severe parasitic infection of the heart—the organ most often affected in
individuals with chronic T. cruzi infection. Symptoms of heart impairment can include thinning of the
ventricular walls, biventricular enlargement, apical aneurysms, and mural thrombi. Widespread destruction of myocardial cells, diffuse fibrosis, edema, lymphocyte infiltration of the myocardium, and scarring of the conduction system are often apparent, but parasites are difficult to find in myocardial tissue
by conventional histologic methods.3,48,49
Chronic Chagas patients exert heave tolls to the public health system because of the resultant morbidity and required hospitalization and treatments. They are a burden on social security because of forced
early retirement and may become marginalized because of their apparent inability to work.42
It is important to distinguish between oral transmission, direct vector-borne transmission, and other
transmission routes when cases are diagnosed; thus, appropriate measures can be implemented to prevent further infections. Various epidemiological, clinical, and laboratory characteristics can be used as
indicators of acute Chagas disease acquired via oral route of infection. These include
1. Acute infection with the absence of cutaneous indurations (chagomas) or Romaña’s sign, having excluded transmission via the transplacental route, organ transplantation, or by accidental
exposure during processing of fresh meat, contact with secretions of infected animals, laboratory accidents, etc.
2. Fever of unknown origin (FUO) in several persons requiring medical attention
Biology of Foodborne Parasites
3. Simultaneous onset of acute symptoms in families or groups of people with a common history
of meals or consumption of artisan beverages
4. Occurrence of specific anti–T. cruzi IgM in several persons, related or not
5. Parasites isolated from infected persons, having the same lineage and clone, based on a similar
nucleotide sequence of hypervariable regions of genes
6. Demonstration of infected vectors and/or reservoirs in the vicinity of where food or beverages
have been processed9
13.7 Treatment and Prevention
The drugs available for treating Chagas disease include only nifurtimox and benznidazole. They are
somewhat effective during the acute phase—antitrypanosomal treatment. Both drugs are extremely
toxic; such therapy is not recommended during pregnancy, except for the most acute and severe
cases. The drug of choice is currently benznidazole. Dosages vary according to age and weight of
patients: for adults, 5 mg/kg/day is recommended; for children, 5–10 mg/kg/day; and for infants,
10 mg/kg/day for 60 days. For adults weighing over 60 kg, the maximum recommended dose of
benznidazole (300 mg/day) should be used for more than 60 days in order to achieve the total dose
needed for patient weight.3,9,24
Given the social and environmental importance of Chagas disease and the great impact it has in the
Americas, disease prevention requires the following: elimination of hematophagous hemipteran vectors through biological control methods or insecticides with long half-life (such as pyrethroids), better
hygiene and housing, cleaning areas around the houses, strict control and screening of donated organs,
serological screening, prenatal follow-up, breastfeeding follow-up, health professional compliance with
laboratory biosafety guidelines, environmental education to fight deforestation and settlements in wild
areas, prevention of domiciliation of vectors and reservoirs, and hygienic food handling and processing
in areas at risk of T. cruzi transmission.
1. Chagas, C. J. R., Nova tripanozomiaze humana. Estudos sobre a morfolojia e o ciclo evolutivo do
Schizotrypanum cruzi n.gen. n.sp., ajente etiolojico de nova entidade morbida do homem. Mem. Inst.
Oswaldo Cruz, 1, 159–218, 1909.
2. Lana, M., Tafuri, W. L., Trypanosoma cruzi e Doença de Chagas. In: Neves, D. P. et al. (Eds.),
Parasitologia Humana, Atheneu, São Paulo, pp. 85–108, 2005.
3. Rassi, A. Jr., Rassi, A., Marcondes de Rezende, J., American trypanosomiasis (Chagas disease). Infect.
Dis. Clin. North. Am., 26(2), 275–91, 2012.
4. WHO. World Health Organization. Chagas disease. Second report of the WHO Expert Committee,
Technical report series N° 905, Geneva, Switzerland, 2002.
5. WHO. World Health Organization. Chagas disease (American trypanosomiasis), Fact sheet N° 340,
2013. Available at: (Accessed February
17, 2014).
6. Dias, J. C. P., O desafio da doença de Chagas nos centros urbanos. Rev. Soc. Brasil. Med. Trop., 32(Suppl. II),
45–48, 1999.
7. Dias, J. C. P. et al., General situation and perspectives of Chagas’ disease in Northeastern Region, Brazil.
Cad. Saúde Pub., 16(1), 13–34, 2000.
8. Camandaroba, E. L., Pinheiro Lima, C. M., and Andrade, S. G., Oral transmission of Chagas’ disease:
Importance of Trypanosoma cruzi biodeme in the intragastric experimental infection. Rev. Inst. Med.
Trop., 44(2), 97–103, 2002.
9. Pereira, K. S. et al., Chagas’ disease as a foodborne illness. J. Food Protect., 72(2), 441–446, 2009.
10. Ribeiro, R. D., Rissato Garcia, T. A., and Bonomo, W. C., Contribuição para o estudo dos mecanismos de
transmissão do agente etiológico da doença de Chagas. Rev. Saúde Pub., 21(1), 51–54, 1987.
Trypanosoma cruzi
11. Forattini, O. P. et al., Nota sobre caso autóctone de tripanossomíase americana no litoral sul do Estado de
São Paulo, Brasil. Rev. Saúde Pub., 14(1), 143–149, 1980.
12. Dias, J. C. P., Notas sobre o Trypanosoma cruzi e suas características bio-ecológicas, como agente de
enfermidades transmitidas por alimentos. Rev. Soc. Bras. Med. Trop., 39(4), 370–375, 2006.
13. Bogitsh, B. J., Carter, C. E., and Oeltmann, T. N., Blood and tissue protozoa I: Hemoflagellates. Human
Parasitol., 85–113, 2005.
14. Brener, Z., Trypanosoma cruzi: Morfologia e ciclo evolutivo. In: Dias, J. C. P. and Coura, J. R. (Eds.),
Clínica e terapêutica da doença de Chagas: uma abordagem prática para o clínico geral, Fiocruz, Rio
de Janeiro, pp. 25–31, 1997.
15. Hamilton, P. B. and Stevens, J. R., Classification and phylogeny of trypanosoma cruzi. In: Telleria, J. and
Tibayrenc, M. (Eds.), American Trypanosomiasis Chagas Disease: One Hundred Years of Research,
Elsevier, London, U.K., Burlington, MA, pp. 321–338, 2010.
16. Kelly, J. M., Taylor, M. C., and Wilkinson, S. R., Trypanosoma cruzi. In: Fuchs, J., Podda, M., and
Goethe, J. W. (Eds.), Encyclopedia of Diagnostic Genomics and Proteomics, Marcel Dekker, Inc., New
York, NY, pp. 1297–1301, 2005.
17. Deane, M. P., Lenzi, H. L., and Jansen, A. M., Trypanosoma cruzi: Vertebrate and invertebrate cycles in the
same mammal host, the opossum Didelphis marsupialis. Mem. Inst. Oswaldo Cruz, 79(4), 513–515, 1984.
18. Jansen, A. M. and Deane, M. P., Trypanosoma cruzi infection of mice by ingestion of food contaminated
with material of the anal gland of the opossum Didelphis marsupialis. Reunião sobre Pesquisa Básica em
Doença de Chagas, Caxambu, MG, 39, 1985.
19. Deane, M. P., Lenzi, H. L., and Jansen, A. M., Double development cycle of Trypanosoma Cruzi in the
Opossum. Parasitol. Today, 2, 5, 1986.
20. Costa, J. et al., The epidemiologic importance of Triatoma brasiliensis as a Chagas disease vector in
Brazil: A revision of domiciliary captures during 1993–1999. Mem. Inst. Oswaldo Cruz, 98(4), 443–449,
21. Costa, J. and Lorenzo, M., Biology, diversity and strategies for the monitoring and control of
triatomines—Chagas disease vectors. Mem. Inst. Oswaldo Cruz, 104(Suppl. I), 46–51, 2009.
22. Center for Disease Control and Prevention (CDC) of United States. Chagas disease. 2014. Available
from: (Accessed January 20, 2014).
23. Pereira, K. S. et al., Trypanosoma cruzi. In: Robertson, L. J. and Smith, H. V. (Eds.), Foodborne Protozoan
Parasites, Nova Science Publishers, Inc., New York, NY pp. 189–216, 2012.
24. Coura, J. R. and de Castro, S. L. A critical review on Chagas disease chemotherapy. Mem. Inst. Oswaldo
Cruz, 97(1), 3–24, 2002.
25. Lescure, F. X. et al., Chagas disease: Changes in knowledge and management. Lancet Inf. Dis., 10(8),
556–570, 2010.
26. Schijman, A. G. et al., International study to evaluate PCR methods for detection of Trypanosoma cruzi
DNA in blood samples from Chagas disease patients. PLoS Negl. Trop. Dis., 5(1), e931 1–e931 13, 2011.
27. Marcon, G. E. B. et al., Use of a nested polymerase chain reaction (N-PCR) to detect Trypanosoma cruzi
in blood samples from chronic chagasic patients and patients with doubtful serologies. Diagn. Microbiol.
Infect. Dis., 43(1), 39–43, 2002.
28. Piron, M. et al., Development of a real-time PCR assay for Trypanosoma cruzi detection in blood samples. Acta Trop., 103(3), 195–200, 2007.
29. Qvarnstrom, Y. et al., Sensitive and specific detection of Trypanosoma cruzi DNA in clinical specimens
using a multi-target real-time PCR approach. PLoS Negl. Trop. Dis., 6(7), e1689 1–e1689 8, 2012.
30. Zingales, B., Trypanosoma cruzi: Um parasita, dois parasitas ou vários parasitas da doença de chagas?
Revista de Biologia, 6b, 44–48, 2011.
31. Miles, M. A. et al., Isozymic heterogeneity of Trypanosoma cruzi in the first autochthonous patient with
Chagas disease in Amazonian Brazil. Nature, 272(5656), 819–821, 1978.
32. Miles, M. A. et al., Further enzymic characters of Trypanosoma cruzi and their evaluation for strain identification. Trans. R. Soc. Trop. Med. Hyg., 74(2), 221–237, 1980.
33. Macedo, A. M. et al., Trypanosoma cruzi: Genetic structure of populations and relevance of genetic variability to the pathogenesis of chagas disease. Mem. Inst. Oswaldo Cruz, 99(1), 1–12, 2004.
34. Morel C. et al., Strains and clones of Trypanosoma cruzi can be characterized by pattern of restriction
endonuclease products of kinetoplast DNA minicircles. Proc. Natl. Acad. Sci. USA, 77(11), 6810–6814,
Biology of Foodborne Parasites
35. Tibayrenc M. and Ayala, F. J. 1987. High correlation between isoenzyme classification and kinetoplast
DNA variability in Trypanosoma cruzi. C. R. Acad. Sci. III, 304(4), 89–92, 1987.
36. Souto, R. P. et al., DNA markers define two major phylogenetic lineages of Trypanosoma cruzi. Mol.
Biochem. Parasitol., 83(2), 141–152, 1996.
37. El-Sayed, N. M. et al., The genome sequence of Trypanosoma cruzi, etiologic agent of chagas disease.
Science, 309(5733), 409–415, 2005.
38. Moncayo, A., Chagas disease: Current epidemiological trends after the interruption of vectorial and
transfusional transmission in the Southern Cone countries. Mem. Inst. Oswaldo Cruz, 98(5), 577–591,
39. Coura, J. R. and Dias, J. C. P., Epidemiology, control and surveillance of Chagas disease—100 years after
its discovery. Mem. Inst. Oswaldo Cruz, 104(Suppl. I), 31–40, 2009.
40. Schmunis, G. A., Epidemiology of Chagas disease in non-endemic countries: The role of international
migration. Mem. Inst. Oswaldo Cruz, 102(Suppl. I), 75–85, 2007.
41. PANAFTOSA. Consulta técnica em epidemiologia, prevenção e manejo da transmissão da doença de
Chagas como doença transmitida por alimentos. Rev. Soc. Brasil. Med. Trop., 39(5), 512–514, 2006.
42. Moncayo, A. and Silveira, A. C., Current epidemiological trends for Chagas disease in Latin America and
future challenges in epidemiology, surveillance and health policy. Mem. Inst. Oswaldo Cruz, 104(Suppl.
I), 17–30, 2009.
43. Shikanai-Yasuda M. A., Carvalho, N. B., Oral transmission of Chagas disease. Clin. Infect. Dis., 54(6),
845–852, 2012.
44. Roque A. L. et al., Trypanosoma cruzi transmission cycle among wild and domestic mammals in three
areas of orally transmitted Chagas disease outbreaks. Am. J. Trop. Med. Hyg., 79(5), 742–749, 2008.
45. Coura, J. R. Tripanosomose, doença de Chagas. Ciência Cultura, 55(1), 30–33, 2003.
46. Britto, C. C. Usefulness of PCR-based assays to assess drug efficacy in Chagas disease chemotherapy:
Value and limitations. Mem. Inst. Oswaldo Cruz, 104(1), 122–135, 2009.
47. Lopes, E. R. and Chapadeiro, E., Pathogenesis of American Trypanosomiasis. In: Maudin, I., Holmes, P.,
and Miles, M. A. (Eds.), The Trypanosomes, CABI International, London, U.K., pp. 303–330, 2004.
48. Rassi, A. Jr., Rassi, A., and Little, W. C., Chagas’ heart disease. Clin. Cardiol., 23(12), 883–889, 2000.
49. Hidron, A. et al., Cardiac involvement with parasitic infections. Clin. Microbiol. Rev., 23(2), 324–349,
Section III
Important Foodborne Helminths
Santhosh Puthiyakunnon and Xiaoguang Chen
14.1 Introduction................................................................................................................................... 235
14.2 Morphology and Classification..................................................................................................... 237
14.3 Biology and Life Cycle................................................................................................................. 238
14.4 Genetics and Genomics................................................................................................................. 239
14.5 Diagnosis....................................................................................................................................... 241
14.6 Epidemiology and Molecular Epidemiology................................................................................ 243
14.7 Pathogenesis and Clinical Features.............................................................................................. 247
14.8 Treatment and Prevention............................................................................................................. 248
14.9 Conclusion..................................................................................................................................... 250
References............................................................................................................................................... 250
14.1 Introduction
Angiostrongylus is a parasitic nematode that can cause severe gastrointestinal or central nervous system
(CNS) disease in humans, depending on the species. Angiostrongylus cantonensis, which is also known
as the rat lungworm, causes eosinophilic meningitis and is prevalent in Southeast Asia and tropical
Pacific Islands. Angiostrongylus costaricensis is another rat parasite causing the human disease, eosinophilic gastroenteritis, and is reported from Costa Rica in and the Caribbean. A. cantonensis was first
discovered in the pulmonary arteries and hearts of domestic rats in Guangzhou (Canton), China, by
Chen1 in 1935. The first human case of angiostrongyliasis was reported by Nomura and Lin2 in Taiwan
in 1945 in a 15-year-old boy, who had 10 worms in his cerebrospinal fluid (CSF) and died as a result of
the infection.3 Since then, many human cases have been reported and infections have been identified in
other areas, including Africa, Caribbean, and the United States.
Humans are accidental hosts of this zoonotic pathogen and may become infected through ingestion of
larvae in raw or undercooked snails or other vectors or contaminated water and vegetables. The larvae
are then transported via the blood to the CNS, where they are the most common cause of eosinophilic
meningitis, a condition that can lead to death or permanent brain and nerve damage.4 Angiostrongyliasis
is an infection of increasing public health importance as globalization has resulted in increased crossborder travel/migration, contributing to the geographic spread of the disease. Major outbreaks of the
disease have been reported in endemic regions, especially mainland China, Taiwan, and the United
States.5 Several major outbreaks of human A. cantonensis infection have been reported in mainland
China in recent years.5
By 2012, about 3161 cases of human angiostrongyliasis had been documented globally (Table 14.1).
However, there are, no doubt, many more cases unreported due to the lack of awareness of this parasite
within the medical community. Presently, however, A. cantonensis has spread from its traditional endemic
regions to the United States, the Caribbean islands, Brazil, Australia, and some areas of Africa where
Biology of Foodborne Parasites
TABLE 14.1
Cases of Human Angiostrongyliasis Reported until 2012
China (including Taiwan and Hong Kong)
Tahiti, French Polynesia
United States
New Caledonia
Réunion Island, France
Sri Lanka
Costa Rica
Côte d’Ivoire
New Zealand
Papua New Guinea
United Kingdom
Total (including 260 new cases since 2010)
Cases (%)
1371 (43.30)
1042 (32.93)
256 (80.91)
134 (4.23)
122 (3.85)
72 (2.27)
63 (1.99)
24 (0.76)
19 (0.60)
11 (0.35)
8 (0.25)
6 (0.18)
6 (0.18)
5 (0.16)
3 (0.09)
2 (0.06)
2 (0.06)
2 (0.06)
2 (0.06)
2 (0.06)
1 (0.03)
1 (0.03)
1 (0.03)
1 (0.03)
1 (0.03)
1 (0.03)
1 (0.03)
1 (0.03)
1 (0.03)
foci of A. cantonensis have been discovered and a number of cases have been reported.6–8 Moreover,
sporadic cases have been reported in travelers after returning from Pacific and Caribbean islands.
When Nomura and Lin first identified this parasite in the CSF, they called the parasite
Haemostrongylus ratti and noted that raw food eaten by the patient may have been contaminated by
rats.2 In 1955, Mackerras and Sandars9 identified the life cycle of the worm in rats, defining snails and
slugs as the intermediate host and noting the path of transmission through the blood, brain, and lungs
in rats. In 1961, an epidemiological study of eosinophilic meningitis in humans hypothesized that the
parasite causing these infections was carried by fish.3 However, Alicata noted that raw fish was consumed by large numbers of people in Hawaii without apparent consequences, and patients presenting
with meningitis symptoms had a history of eating raw snails or prawns in the weeks before presenting
with symptoms. This observation along with epidemiologic investigations and autopsy of infected
brains confirmed A. cantonensis infection in humans as the cause of the majority of eosinophilic meningitis cases in Southeast Asia and the Pacific Islands.10 Since then, this zoonotic parasitic infection is
commonly called “Alicata’s disease.”11
In recent years, the parasite has been shown to be proliferating at an alarming rate due to modern food
consumption trends and global transportation of food products. Scientists are calling for a more thorough
study of the epidemiology of A. cantonensis, stricter food safety policies, and improvements in education
on proper consumption of products commonly infested/contaminated by the parasite.12
14.2 Morphology and Classification
A. cantonensis is a helminth parasite classified in the phylum Nematoda, class Secernentea, order
Strongylida, and superfamily Metastrongyloidea and is commonly referred as the “rat lungworm.” There
are obvious differences between male and female A. cantonensis. The males are 15.9–19 mm in length,
while the females can grow 21–25 mm. Females are easily distinguished from males by the noticeable
barber-pole appearance in their bodies (Figure 14.1). This is actually the interweaving of the intestine
and uterine tubules. Females have a vulva, which is located 0.2 mm in front of the anus.13,14
A. cantonensis is a flimsy and slender cylindrical worm and has a cuticle with three main outer layers
made of collagen and other compounds. The outer layers are noncellular and are secreted by the epidermis. The cuticle layer protects nematodes from enzymes in the digestive tracts of animals. The worms
molt four times during the entire life cycle, the first two times in intermediate hosts, and the second two
times in definitive hosts. As a member of the Secernentea, A. cantonensis has a specialized tubular excretory system with three canals.13,15 The canals are arranged to form an “H” shape. The bursa, a structure
used to clasp females when copulating, is small, and the dorsal lobe is not present. The males have long
and slender spicules, which are almost equal in length and form.14
Females may produce a pheromone to attract males. The male coils around a female with its curved
area over the female genital pore. The gubernaculum, made of cuticle tissue, guides spicules, which
extend through the cloaca and anus. Males use spicules to hold the females during copulation. Nematode
sperm are amoeboid like and lack flagella.13–15
A. cantonensis is able to swim intermittently. The worms are usually only able to move effectively
when the pseudocoel is filled with fluid and hypertonic to the surrounding media.14 These parasitic
nematodes have phasmids, which are unicellular glands and likely function as chemoreceptors. The
other structures include papillae and setae, which are the main sense organs and help the worms to
detect motion (mechanoreceptors), respectively.14,15 A. cantonensis has a simple mouth and no buccal
cavity. The pharyngeal glands and intestinal epithelium produce digestive enzymes and feed on the body
fluids of its hosts. Extracellular digestion begins within the intestine, and the digestive cycle is completed
FIGURE 14.1 Morphology of A. cantonensis. (a) A. cantonensis third-stage (L3), infective larva recovered from a slug.
Image captured under differential interference contrast (DIC) microscopy. (b) Angiostrongylus adult worm recovered from
vitreous humor of a human patient. The bursa is one indication that this is a male worm. (c) Adult male, note copulatory
bursa at posterior of male. (d) Adult female with characteristic “barber-pole” spirals.
Biology of Foodborne Parasites
14.3 Biology and Life Cycle
A. cantonensis is not specific in using either definitive or intermediate hosts. The requirement is that
the intermediate host must be an invertebrate, while the definitive is a terrestrial mammal (Figure 14.2).
Paratenic hosts (an intermediate host in which no development of the parasite occurs) can be either
invertebrate or vertebrate. The definitive hosts for A. cantonensis are usually rodents from the genus
Rattus, with some main ones being Rattus norvegicus and Rattus rattus. A. cantonensis can also
Larvae mature, lay eggs,
and hatch first-stage
larvae in lungs
First-stage larvae passed
in faeces
Third-stage larvae
Vegetables contaminated
with third-stage larvae
Larvae reach CNS,
cause eosinophilic
Third-stage larvae
in snails or slugs
Third-stage larvae in land crabs,
frogs, freshwater prawns, monitor
lizards, and planarians
Larvae enter
in intestine
FIGURE 14.2 The life cycle of A. cantonensis. (a) The adult worms develop to sexual maturity and lay eggs in the pulmonary
arteries. (b) The eggs are hatched into first-stage larvae (the juveniles), which are swallowed and are excreted out with the feces.
(c) The larvae in feces are swallowed by intermediate host mollusks (snails or slugs) and develop into third-stage (infective)
larvae. (d) The third-stage larvae are then transmitted to the paratenic hosts such as shrimps, land crabs, predacious land planarians, and monitor lizards. (e) Humans occasionally acquire A. cantonensis when they eat snails and slugs and sometimes land
crabs, frogs, freshwater shrimps, monitor lizards, or vegetables, which contain the infective larvae. (f) The larvae are digested
from tissues and enter the bloodstream in the intestine. (g) The larvae finally reach the CNS and cause eosinophilic meningitis or
move to the eye chamber and cause ocular angiostrongyliasis. (Adapted from Wang, Q.P. et al., Lancet Infect. Dis., 8, 621, 2008.)
survive in humans and monkeys. Two cases of monkeys dying from complications of eosinophilic
meningoencephalitis in zoos were attributed to their contact with intermediate hosts snails.13,14,16,17
Once inside the intestine of the definitive host, the larvae go through obligatory migration through the
CNS via the bloodstream to the brain and spinal cord. They leave the capillaries and wander randomly
through the tissues, during which they develop into the fifth-stage larvae.14 After the larvae reach the surface of the brain or spinal cord, they penetrate the veins to reenter the circulatory system. The larvae end
up at the pulmonary arteries where they mature into adults in about 6 weeks. However, some larvae wander
to other places in the body and mature there. Common places where works are found include the CNS, the
meninges, and the eyes.13,17 The larvae are also found circulating in the blood, spinal fluid, CSF, or in the
blood vessels of the brain and the meninges.13,14
Humans are accidental hosts; the parasite cannot reproduce in humans, and therefore, humans do not
contribute to the A. cantonensis life cycle. In humans, the circulating larvae migrate to the meninges
but do not move on to the lungs. Sometimes, the larvae will develop into the adult form in the brain and
CSF, but they quickly die, inciting the inflammatory reaction that causes the symptoms of infection.18
Intermediate hosts of this nematode are mollusks, but crustaceans (prawns and land crabs), predacious
land planarians (flatworms in the genus Platydemus), frogs, and monitor lizards can serve as paratenic
hosts. Adult worms of A. cantonensis commonly live in the pulmonary arteries or rarely in the heart
of domestic rats.19 Rats, as definitive hosts, are infected with A. cantonensis after ingesting third-stage
larvae. The larvae migrate to the CNS, where they become the fourth- and fifth-stage larvae via two
molts and finally develop to adult worms.20 However, the worms undergo sexual maturity and lay eggs
in pulmonary arteries. Eggs hatch into first-stage larvae, which migrate up the bronchial tree, are swallowed and are excreted out with the feces.21
The larvae penetrate, or are ingested by, an intermediate host (snail or slug). After two molts, the thirdstage larvae are produced, which are infective to mammalian hosts. When the mollusk is ingested by
the definitive host, the third-stage larvae migrate to the brain where they develop into young adults. The
young adults return to the venous system and then the pulmonary arteries where they become sexually
mature.21 Paratenic hosts that ingest the infected snails can carry the third-stage larvae that can resume
their development when the paratenic host is ingested by a definitive host.19 Humans can acquire the
infection by eating raw or undercooked snails or slugs infected with the parasite; they may also acquire
the infection by eating raw produce such as lettuce and watercress that contains a small snail or slug or
part of one.5,19 The disease can also be acquired by ingestion of contaminated or infected paratenic hosts
(crabs, freshwater shrimps, frog, lizards, and predacious land planarians). In humans, juvenile worms
migrate to the brain, or rarely in the lungs, where the worms ultimately die (Figure 14.2).
The life cycle of A. (Parastrongylus) costaricensis is similar, except that the adult worms reside in the
arterioles of the ileocecal area of the definitive host. In humans, A. costaricensis often reaches sexual
maturity and release eggs into the intestinal tissues. The eggs and larvae degenerate and cause intense
local inflammatory reactions and do not appear to be shed in the stool.5,19,22
14.4 Genetics and Genomics
The genome of A. cantonensis has not been fully sequenced, but some studies have been conducted
on the genes and proteins of this parasite. A cDNA library of A. cantonensis fourth-stage larvae that
were isolated from the brain of artificially infected mice was constructed, and 1200 clones have been
sequenced.19 The fourth-stage larvae are similar to those found in human who are infected with this
parasite. Bioinformatics assay revealed the presence of 378 cDNA clusters, of which 168 contained
open reading frames encoding proteins containing an average of 238 amino acids. Characterization
of these encoded proteins by gene ontology analysis showed enrichment in proteins with binding and
catalytic activity.23 In addition, a total of 1277 expressed sequence tags (ESTs) of A. cantonensis were
randomly downloaded from NCBI and analyzed. Of these, 695 ESTs were grouped into 13 categories based on fucntions.24 In addition, some proteins of A. cantonensis such as cystatin,25 galectin,26
and γ-butyrobetaine hydroxylase27 have been cloned and expressed, and their functions have been
Biology of Foodborne Parasites
In a recent study, a novel gene encoding galectin-10 (AcGal-10), which plays an important role in host–
parasite interactions, was characterized from a cDNA library of A. cantonensis and its biological role in
the parasite examined.28 Analysis of recombinant AcGal-10 revealed that it plays a role in the activation
of the microglia, and this finding suggests that AcGal-10 may be an important molecule related to infection of A. cantonensis.28
An activation-associated secreted protein (ASP) has also been found to play a role in the pathogenesis
of human angiostrongyliasis. A cDNA sequence encoding a putative two-domain ASP was obtained
from an A. cantonensis fourth-stage larva cDNA library, designated as AgASP, which contains an open
reading frame encoding 424 amino acids, with the first 19 residues being a putative secretion signal.29
The expression pattern of this protein was investigated using quantitative real-time polymerase chain
reaction (PCR) (qRT-PCR) and Western blotting and revealed that this protein was highly expressed
in the brain-stage larvae (Lbr) and was found in the excretory/secretory (ES) products of this stage.
Immunofluorescence showed that it existed in the lumen of the brain-stage larvae.
A phylogenetic analysis was conducted on isolates of A. cantonensis from different geographical locations in Brazil, using mitochondrial cytochrome c oxidase subunit I (COI) gene sequences.30 A distinct
monophyletic clade was identified that included all isolates of A. cantonensis from Brazil and Asia
based on eight distinct haplotypes (ac1, ac2, ac3, ac4, ac5, ac6, ac7, and ac8) from a previous study.
Interestingly, the Brazilian haplotype ac5 clustered with isolates from Japan, and the Brazilian haplotype ac8 from the Brazilian states of Rio de Janeiro, Sao Paulo, Pará, and Pernambuco formed a distinct
clade. A divergent Brazilian haplotype, which was named ac9, was closely related to the Chinese haplotype ac6 and the Japanese haplotype ac7. These findings support the hypothesis that the appearance of
A. ­cantonensis in Brazil is likely a result of multiple introductions of parasite-carrying rats, transported
on ships due to active commerce with Africa and Asia during the European colonization period. The
rapid spread of the intermediate host, Achatina fulica, also seems to have contributed to the dispersal of
this parasite and the infection of the definitive host in different Brazilian regions.30
Thaenkham et al.31 from Thailand examined the genetic variation and haplotype relationships using
RAPD-PCR on parasites obtained from eight different geographic areas of Thailand. Based on eight
primers, high levels of genetic diversity and low levels of gene flow among populations were found.
A. cantonensis in Thailand could be divided into two groups with statistically significant genetic differentiation of the two populations using distance and neighbor-joining dendrogram methods,31 but further
elucidation using other genotypic methods is required to confirm this.
A cDNA encoding a protein disulfide isomerase (AcPDI) was cloned and characterized from an
A. cantonensis cDNA library from the fourth-stage larvae (L4) and exhibited high identity to the homologues from other species.32 qRT-PCR performed on the third- and fourth-stage larvae, and adult stages
of A. cantonensis revealed that the AcPDI mRNA was expressed at a significantly higher level in female
adult worms. Immunohistochemical studies further confirmed that AcPDI expression was specifically
localized in the tegument and uterus wall of female adult worms. Collectively, these results implied
that AcPDI may be a female-enriched protein and is associated with the reproductive development of
A. ­cantonensis and cuticle formation.32 Another novel gene encoding a cathepsin B–like cysteine protease (AcCBL1) was recently identified from an A. cantonensis L4 cDNA library, and its biological role
in the parasite was characterized.33 The identification of a “hemoglobinase motif” confirmed its possible
role in the degradation of hemoglobin and its importance in parasite nutrition uptake.
Chang et al. conducted a transcriptomic study of pepsin-activated infective larvae of A. cantonensis;
1496 ESTs were generated from a cDNA library and clustered into 161 contigs and 757 singletons.34
Among these unigenes, 54.5% had significant sequence homology to known proteins and most of them
were abundantly expressed transcripts like cathepsin B–like cysteine proteases 1 and 2, metalloprotease 1, metalloprotease 1 precursor, and extracellular superoxide dismutase. Moreover, 280 clusters were
mapped to 158 KEGG (Kyoto Encyclopedia of Genes and Genomes—
html) pathways and 134 had unique EC (reference pathway) numbers. These findings suggest that treatment with pepsin–HCl not only digests the tissues of the snail host but also activates the infective larvae.
Recently, a genetic sequencing study was carried out to determine and characterize microRNAs (miRNAs) of female and male adults of A. cantonensis using Solexa deep sequencing.35 miRNAs are endogenous small noncoding RNAs that play crucial roles in gene expression regulation, cellular function and
defense, homeostasis, and pathogenesis of many known parasites. A total of 8,861,260 and 10,957,957
high-quality reads with 20 and 23 conserved miRNAs were obtained in females and males, respectively.
No new miRNA sequence was found. This report of miRNA profiles in A. cantonensis may be useful in
studying gene regulation in A. cantonensis in future.
14.5 Diagnosis
In humans, Angiostrongylus is the most common cause of eosinophilic meningitis or meningoencephalitis.5 Infection first presents with severe abdominal pain, nausea, vomiting, and weakness, which gradually lessens and progresses to fever, and then to CNS symptoms and severe headache and stiffness of the
neck. Worms can also invade the eye chamber resulting in visual impairment, pain, keratitis, and retinal
edema. A. cantonensis infections result in eosinophilic meningitis, while A. costaricensis infections
cause eosinophilic enteritis.
Diagnosis of human A. cantonensis is based on clinical features as well as laboratory findings. The
diagnosis of A. cantonensis meningitis is suggested by the presence of typical features of the disease,
eosinophilic pleocytosis of the CSF, and a history of consumption of food likely to contain infective larvae in endemic areas. Important clues that could lead to the diagnosis of infection in nonendemic areas
are a history of travel to where the parasite is known to be found and ingestion of raw or undercooked
snails, slugs, or possibly paratenic hosts (such as frogs, freshwater shrimp, or land crabs) in these areas.
Laboratory findings in blood and CSF, brain imaging, serological assays, and molecular tests are also
used in the diagnosis of human angiostrongyliasis.
The recovery of A. cantonensis from CSF or the ocular chamber confirms human angiostrongyliasis but
is difficult in practice. A spinal tap, or a sample of CSF, must be taken to search for A. cantonensis worms
or larvae. Placing a patient in a sitting position for 1 h before a lumbar puncture has been shown to increase
the yield of larvae in the CSF.36 In many cases, A. cantonensis is undetectable in the CSF as it can adhere
to the meninges. The worm in the eye can be detected by slit lamp or funduscopic examination, and it is
recommended that this examination be performed in any individual presenting with a history of eating
raw intermediate hosts and visual loss, with or without eosinophilic meningitis.37 Because the frequency of
detecting these worms in patients is very low, the organism is rarely detected by microscopy. Adult worms
were only recovered from 28 (11%) of 259 human cases in Taiwan38 and 10 (2%) of 484 cases in Thailand.39
During an active infection, both intracranial pressure and eosinophil blood counts rise, and the
latter may increase up to 35% of total white blood cell population in peripheral blood (normal range
­0.5%–5%).36 Eosinophilic meningitis is usually confirmed by detecting a higher percentage of eosinophils in the blood (>5%) or in CSF (>10%).36,38,40,41 It is important to note, however, that eosinophilia in
the CSF and peripheral blood may not be present on initial presentation or in late stages of infection.5
The chemical analysis of the CSF typically resembles the findings in aseptic meningitis with slightly
elevated protein levels, normal glucose levels, and negative bacterial cultures.5 The presence of a significantly decreased glucose on CSF analysis is an indicator of severe meningoencephalitis and may indicate
a poor prognosis. A. costaricensis is usually found in the intestine (especially the ileocecal region) and
can cause abdominal pain, fever, nausea, and vomiting. As with A. cantonensis, elevated blood eosinophils are an important diagnostic clue.
Neuroimaging studies such as MRI and CT have been used to detect damage in brain for differential
diagnosis of A. cantonensis from other parasites.42–44 MRI and CT reveal focal lesions in the CNS and
can be useful for the differential diagnosis of the disease from other parasitic diseases such as cysticercosis, paragonimiasis, gnathostomiasis, and schistosomiasis.38,42–47 MRI findings in A. cantonensis
infection include abnormal enhancing lesions in the brain, especially hyperintense T2 signal lesions,
which are different from the hemorrhagic lesions in Gnathostoma spinigerum infections.47,48 CT scan of
the brain can be normal or can show nonspecific findings, including cerebral edema and infarction, with
areas of enhancement due to vasculitis and meningitis, ventricular dilatation with hydrocephalus,49 and
diffuse meningeal enhancing ring or disc lesions, resembling a tuberculoma.45,48 Recently, Shih et al.
reported that chest radiographs may show dense opacities with ill-defined margins and segmental distribution in the lower lung fields, presumably due to either vascular thrombosis or larval track reactions.49,50
Biology of Foodborne Parasites
Immunological assays are important for the diagnosis of both A. cantonensis and A. ­costaricensis
infections since these parasites cannot be isolated from fecal matter and are rarely found in CSF
­samples. However, although serologic tests have been developed to effectively diagnose and manage
A. ­cantonensis infection, they are yet to be widely available for commercial use. Both A. costaricensis
and A. cantonensis share common antigenic epitopes, which elicit antibodies that recognize proteins
present in both species. Various immunologic detection methods including ELISAs have been developed
to detect the antigens of or antibodies against A. cantonensis in serum or CSF.51–58 These have been
shown to be both specific and sensitive (approaching 100%), but the instability of reagents and the need
for sophisticated equipment are among the factors limiting their use in the field. A dot-blot ELISA has
shown promise in fulfilling the requirements of an economic and simple field test.51,52,55
As with any other nematode infection, the key to specific diagnosis is the use of an appropriate antigen.
In the past several years, progress has been made in the identification of the antigens that are specifically
diagnostic for human A. cantonensis.53–55,59–64 One study isolated and characterized an immunogenic
31 kDa A. cantonensis protein present in crude adult worm extracts and reported that reactivity to the
31 kDa protein may represent antibody recognition of more than one protein.53
Recently, we analyzed the diagnostic value of larval ES antigens (LESA) in A. cantonensis infection and compared them with adult worm antigens (AWAs) using SDS-PAGE and Western blotting.54
The serum levels of IgG and IgM antibodies to LESA increased at the beginning of infection in mice,
reaching a peak on day 5 after infection and decreased on day 10. Compared with AWA-ELISA, LESAELISA showed a lower seropositive ratio in suspected patients with A. cantonensis, with also a lower
cross-­positive ratio in patients with schistosomiasis and Clonorchis sinensis. Although LESA had a
slightly lower positive ratio than AWA, LESA had a higher specificity for detecting serum antibodies in
suspected cases of A. cantonensis infection and, therefore, shows a potential for the diagnosis of angiostrongyliasis especially in the early stage and in current infection.
Recently, four murine monoclonal antibodies (MAbs) were generated against the ES products of
A. cantonensis adult worms, and a double antibody sandwich ELISA and gold immunochromatography
assay (GICA) were developed using two MAbs (12D5 and 21B7) to detect the circulating antigen (CAg)
in the sera of rats infected with A. cantonensis and angiostrongyliasis patients, respectively. Results from
the sandwich ELISA and GICA showed high specificity (100%), indicating that these rapid and simple
tests might be useful in early detection of angiostrongyliasis cases.57
Using this chromatography assay, a seroepidemiological survey of 1730 blood samples was conducted
in China during 2009–2010 to determine the extent of CAgs of A. cantonensis in the Chinese population.59 The overall seroprevalence in the “occupational group” (i.e., those involved in aquaculture or
processing of the snails Ac. fulica and Pomacea canaliculata) was 7.4%, which was significantly higher
than that in the “general group” (general adult population) (0.8%). These results suggest that angiostrongyliasis represents a significant zoonotic disease in China, and the strengthening of food safety is needed
for the control of this foodborne disease.
In the last decade, various DNA-based molecular diagnostic methods have become available for
detecting nucleic acids of pathogenic parasites in biological materials such as blood, stool, urine, cutaneous scrapings, and secretions, including CSF.65 Recently, the mRNA sequence encoding the 66 kDa
native protein from A. cantonensis adult worms has been described and used as the basis for developing
a PCR protocol for the specific identification of species of the genus Angiostrongylus.66–68 A single-step
PCR A. cantonensis DNA in CSF samples from 4 of 10 serologically confirmed neuroangiostrongyliasis
cases. In addition, an immuno-PCR has been developed for detecting A. cantonensis in CSF and tissue.60 A multiplex PCR method has recently been developed for the detection of A. cantonensis larvae in
P. canaliculata (apple snail) and exhibited 93.8% sensitivity and 80.6% specificity.69
A loop-mediated isothermal amplification (LAMP) method for the specific detection of A. cantonensis in the snail Ac. fulica has also been developed.70 Primers for LAMP were designed based on the first
internal transcribed spacer (ITS-1) of nuclear ribosomal DNA (rDNA) of A. cantonensis. The assay was
rapid, inexpensive, specific for A. cantonensis, and 10 times more sensitive than the conventional PCR
assay. A species-specific internal transcribed spacer 1–based TaqMan assay for A. cantonensis has been
developed,71 and more recently, a quantitative PCR (qPCR) assay was developed using the same primers
and probe to provide a reliable, relative measure of parasite load.72
A protein microarray for the rapid screening of patients suspected of infection with cysticercosis, trichinellosis, paragonimiasis, sparganosis, and angiostrongyliasis was recently developed.73 Semipurified
antigens from Cysticercus cellulosae, A. cantonensis, Paragonimus westermani, Trichinella spiralis,
and Spirometra sp. were screened against sera from 365 human cases of helminthiasis and 80 healthy
individuals, and the results were compared with ELISA. The protein microarray had specificity (97%–
100%) and sensitivity (85.7%–92.1%) similar to the ELISA (97.7%–100% v 82%–92.1%).73 As multiparasitic infections are very common, this protein microarray provides a sensitive, high-throughput
technique for the simultaneous detection of multiple foodborne helminths.73
14.6 Epidemiology and Molecular Epidemiology
A. cantonensis has become an important emerging pathogen worldwide due to frequent outbreaks and
increased reports of sporadic cases in new endemic areas. Previously, human cases were exclusively
reported in known endemic areas like the Pacific Islands and Southeast Asia. However, thousands of
cases of human angiostrongyliasis have been documented worldwide in the last few decades as globalization has allowed it to spread to the other locations including the United States, Caribbean islands,
Brazil, and Australia. Since the first case of human angiostrongyliasis was reported in 1945, more than
3160 cases have been reported in approximately 30 countries until 2012 (Table 14.1). Most of the endemic
areas of this parasitic disease are popular tourist sites. Thus, increasing cross-border travel and tourism
contribute significantly to its broad geographic distribution, along with the growing popularity of eating
exotic foods such as raw and undercooked snails.
The highest number of human cases had been reported in Thailand, which accounts for about 43% of
the total cases reported worldwide. The high rate of the disease in the Thai population is linked to the
custom of eating raw or undercooked snails (Pila spp.) with alcohol, which is especially popular among
young men.74,75 Nearly 70% of Thai patients infected with A. cantonensis were 20–40 years old.39 By
contrast, 65% of the reported 125 cases in Taiwan have been in children under 10 years of age.38
Outbreaks of human infections caused by A. cantonensis linked to contaminated food consumption
have increased significantly in the past decade in mainland China, where more than 650 million people
are estimated to be at risk (Table 14.2). In mainland China, the first case of human angiostrongyliasis
with eosinophilic meningitis was reported in 1984 in Guangdong province.76 Many more outbreaks of
human infection with A. cantonensis have been reported with increasing frequency in other provinces
TABLE 14.2
Outbreaks of Human Angiostrongyliasis Reported in Mainland China since 1997
Regions (Province)
Number of Cases
Biology of Foodborne Parasites
of mainland China in recent years. The largest outbreak was reported in Beijing in 2006 with 160 cases.
The majority of other human A. cantonensis outbreaks had been reported between 1997 and 2011. One
outbreak of human A. cantonensis infection with 65 cases of eosinophilic meningitis was observed in
Wenzhou, Zhejiang province, in 1997.77 An outbreak of five human cases occurred in Liaoning province
in 1999,78 and three outbreaks with a total of 30 cases were reported in Fujian province in 2002.79–81 Four
outbreaks with 101 human cases occurred in Yunnan since last decade.82,83 A recent outbreak of angiostrongyliasis was reported in Dali, Yunnan, after a group of 16 persons consumed food with snails.84
The infectivity rate was 56.3% (9 of 16 patients), and three cases were confirmed by serology (42.9%).
Surveillance in this area revealed that 7.3% of snails (Pomacea spp.) were infected. Angiostrongyliasis
is common among many migrant workers from this area. There was another outbreak of angiostrongyliasis among 17 migrant laborers in Guangning, Guangdong province, and all of them belonged to the
Bai ethnic group from Dali.85 All patients consumed P. canaliculata, and six had meningitis 3–19 days
after consumption of P. canaliculata, and five of their blood samples were positive for antibodies to
A. ­cantonensis. This outbreak highlights the vulnerability of migrants to angiostrongyliasis.
After Thailand and China, the majority of human cases have been reported from the Caribbean islands
and the United States. The first case of human angiostrongyliasis from these countries was reported in
Cuba in 1973,86 followed by several cases reported in Costa Rica and Jamaica.40,87,88 In the United States,
cases have been reported from Hawaii, which is an endemic area.18 The infection is now endemic in
wildlife, and a few human cases have also been reported in areas where the parasite was not originally
endemic, such as New Orleans and Egypt.89 Many cases have been reported in travelers returning from
endemic areas. Of a group of 23 U.S. travelers, 12 developed eosinophilic meningitis after returning
from Jamaica in 2000.40 Additionally, other sporadic cases of human A. cantonensis infection have been
described in travelers returning from the Pacific or Caribbean islands.5
Most species of mollusks are susceptible to and capable of transmitting A. cantonensis (Table 14.3).
The prevalence of infection in terrestrial snails and slugs is higher than in freshwater mollusks, indicated
by a recent national survey conducted in China.90 Analysis of prevalence rates in snail species in China
revealed that 22 of 32 species of wild mollusks (69%) were infected with the parasite.12 Terrestrial and
some aquatic snails are the primary intermediate hosts.12,74 The giant African snail Ac. fulica is the major
source of infection worldwide and has been recorded with the highest rate and intensity of infections, followed by slugs (Vaginulus spp.) and golden apple snail, P. canaliculata, although other snails and slugs
can also be vectors. P. canaliculata, which is native to South America, was imported to Taiwan in 1981
as a food source91,92 and then introduced to mainland China. The infection rate in P. ­canaliculata is very
high in Taiwan (21%), mainland China (42%–69%), and Okinawa, Japan (10%–39%).93
The dispersal of A. cantonensis was associated with the spread of this snail from its native origin in
Africa throughout the Pacific Islands and South Asia.91 The golden apple snail P. canaliculata has a
wide distribution in Asia and has caused great damage to local agricultural systems. Unfortunately, this
snail is also very susceptible to A. cantonensis and has become an important intermediate host in these
regions.94 In Thailand, although Pila snails are frequently eaten by men, these snails are poor vectors and
contain less infective A. cantonensis worms. Consequently, human infections resulting from eating Pila
spp. have milder clinical signs than those caused by eating P. canaliculata.74
In a study in China, P. canaliculata and Ac. fulica were found in 11 and 6 provinces, respectively. Out
of 11,709 P. canaliculata snails examined, 6.8% were infected with A. cantonensis. Of 3549 Ac. fulica
snails examined, 13.4% were infected with A. cantonensis. P. canaliculata has replaced Ac. fulica as
the major source of human infection, playing an important role in the epidemiology of A. cantonensis
in recent outbreaks of human angiostrongyliasis.12 This was confirmed by a recent study demonstrating that P. canaliculata had an average infection rate of 21%, which is significantly higher than that of
Ac. fulica (10%) in Shenzhen, Guangdong province.95 It was thought that the larvae of A. cantonensis
can be released from mollusks into slime fluid and contaminate produce and other objects as they crawl.
However, Liang et al.96 did not detect larvae in body fluids washed from 23 Ac. fulica snails that were
infected with A. cantonensis. Therefore, whether slime fluid plays a role in human infection remains
A study was conducted between 2008 and 2010 to reassess the enzootic angiostrongylus infection
status of rodent definitive hosts, snail intermediate hosts, and local residents in Guangzhou, China.
TABLE 14.3
Various Reported Hosts of Angiostrongylus cantonensis
Host Type
1. Definitive hosts
Wild rodents
Domestic rats
2. Intermediate hosts
Land snails
Giant African snail
Golden apple snail
Freshwater snails
3. Paratenic hosts
Predatory land flatworm
Freshwater prawns
Land crabs
Monitor lizard
Animal Species
a. Rattus norvegicus
b. Rattus rattus
a. Thelidomus aspera
b. Achatina fulica
c. Satsuma mercatoria
d. Acusta despecta
e. Bradybaena brevispina
f. Bradybaena circulus
g. Bradybaena ravida
h. Bradybaena similaris
i. Plectotropis appanata
j. Parmarion martensi
k. Camaena cicatricosa
l. Trichochloritis rufopila
m. Trichochloritis hungerfordianus
n. Cyclophorus spp.
o. Pomacea canaliculata
a. Pila spp.
b. Pomacea canaliculata
c. Cipangopaludina chinensis
d. Bellamya aeruginosa
e. Bellamya quadrata
a. Limax maximus
b. Limax flavus
c. Deroceras laeve
d. Deroceras reticulatum
e. Veronicella alte
f. Laevicaulis alte
g. Sarasinula plebeia
h. Vaginulus yuxjsjs
i. Lehmannia valentiana
j. Philomycus bilineatus
k. Macrochlamys loana
l. Meghimatium bilineatum
Platydemus manokwari
a. Bufo asiaticus
b. Rana catesbeiana
c. Rhacophorus leucomystax
d. Rana limnocharis
e. Hyla aurea
Decapoda spp.
Pleocyemata spp.
Varanus bengalensis
(Continued )
Biology of Foodborne Parasites
TABLE 14.3 (Continued )
Various Reported Hosts of Angiostrongylus cantonensis
Host Type
4. Accidental hosts
Yellow-tailed black cockatoo
Tawny frogmouths
Brushtail possums
Human beings
Animal Species
Calyptorhynchus funereus
Podargus strigoides
Trichosurus vulpecula
Homo sapiens
The average infection rate in rats (R. rattus), Ac. fulica snails, and P. canaliculata snails were 5.35%,
13.96%, and 1.50%, respectively.97 A high seroprevalence was also noticed in the “occupational group”
(those individuals involved in aquaculture or the processing of Ac. fulica and P. canaliculata snails)
compared to general human population. This survey shows that residents who live in Guangzhou, especially those working in certain industries such as agriculture, food processing, and aquaculture, face a
higher risk of infection.
Quantification of the levels of infection in intermediate hosts is critical in determining the extent of
A. cantonensis endemicity in an area where the disease is prevalent. The recent development of a qPCR
assay for quantifying A. cantonensis infections in intermediate hosts72 will facilitate the identification
of high-intensity infections and allow the implementation of more effective targeted rat and slug control
Rats (R. rattus and R. norvegicus) and other rodents are the definitive host and the main reservoir
for A. cantonensis, although other species of rats found in rural and forested areas are also reported to
be natural hosts21,74 (Table 14.3). Rats are necessary for the establishment of A. cantonensis foci in an
area as they act as a continuous source of infection to maintain its life cycle in its endemic area. When
A. cantonensis is identified in rats, the parasite is deemed to be endemic in that region. A study indicating the prevalence of infection in definitive hosts revealed that 11 of 15 wild rodent species in mainland
China were infected with A. cantonensis.12 R. norvegicus is the most frequently identified host with a
generally higher prevalence and intensity of infection compared with other rodents. This was consistent
with a national survey in China that found 32 of 711 rats infected with A. cantonensis.90 Norway rats
(R. norvegicus) were originally native to northern China. Following a series of introductions, the species
found its way to Eastern Europe by the early eighteenth century. By the year 1800, they occurred in every
European countries. Records show the first sighting of R. norvegicus in the New World in the 1770s as
ship stowaways. Today, R. norvegicus (also known as brown rats) can be found on every continent of the
world except Antarctica.98,99
Angiostrongylosis is common in dogs in Australia, and they may be reservoirs of infection for humans.
Between 2002 and 2005, 22 cases of canine neural angiostrongylosis (NA) were reported in Australia.100
CSF evaluations showed that 59 suspected dogs from the same areas (South East Queensland and
Sydney) also had neural symptoms.100 Numerous cases in tawny frogmouths were reported from the
same regions as the affected dogs over the study period.100 These findings strongly suggests the existence of an enzootic cycle of angiostrongyliasis as an outcome of close interrelationship among many
unnatural hosts, known natural hosts, and the common paratenic hosts. Interestingly, A. cantonensis was
also found in nonhuman primate, equine, and canine species.90 A. cantonensis was also discovered in
paratenic host frog species (Hylarana guentheri, Rana limnocharis, and Rana plancyi) and toads (Bufo
­melanostictus) but has not yet been identified in freshwater shrimp, fish, crabs, or planaria in published
studies.12 A. ­cantonensis was not found in any of 652 paratenic hosts collected during a national survey
in China90 that included frogs, shrimps, crabs, toads, and fish.19
Other than the intermediate hosts, some animals are also important for the transmission of this
parasitic nematode. Even though there is little knowledge regarding the prevalence of A. cantonensis
in paratenic hosts,5 some of them play a very crucial role in small outbreaks of angiostrongyliasis.
Predatory land planarians (flat worms), Platydemus manokwari, could represent a very important but
overlooked source of human infection when they are consumed together with contaminated uncooked
vegetables. Four outbreaks of human angiostrongyliasis have been caused by eating contaminated
vegetables or vegetable juice.101 An outbreak of five cases in Taiwan was linked to drinking vegetable
juices in 2001.101
In New Caledonia, 53% of frogs (Hyla aurea) were found to contain the infective larvae.102 Eating
raw frogs has been implicated in human infections in Taiwan, mainland China, and the United States.
In Thailand, 21 (95%) of 22 monitor lizards studied were found to be infected with A. cantonensis,
and more than 18 cases of human angiostrongyliasis in Thailand, Sri Lanka, and India were attributed
to consumption of monitor lizards.103–105 Humans and nonhuman primates can be accidental hosts for
A. cantonensis, but the parasite is unable to complete its development and usually dies in the CNS, causing eosinophilic meningitis or even death. The worms have been reported as the cause of death of captive
primates in the Bahamas, Australia, and the United States.6,106 More recently, A. cantonensis has become
well established in Sydney where it has caused disease in tawny frogmouths, domestic dogs, grey-headed
flying foxes, a brushtail possum, and humans.107 The parasite was originally thought to be restricted to
Queensland where infection has been reported in captive species of birds and animals. It is likely that
the distribution of A. cantonensis will continue to expand, as many species of native and introduced terrestrial mollusks in Australia are suitable intermediate hosts.107
14.7 Pathogenesis and Clinical Features
The incubation period of the parasite in human angiostrongyliasis ranges from 1 day to several months,21
depending on the number of parasites involved. In an outbreak in Beijing, China, the incubation period
for 80% of patients was 7–35 days.108 In an outbreak in Wenzhou, China, clinical symptoms appeared in
62% of patients 6–15 days after infection.109 Humans acquire A. cantonensis after eating intermediate or
paratenic hosts or vegetables that contain the infective larvae (the third stage) of the worm. Once swallowed, the infective larvae are digested from those vectors and invade intestinal tissue, causing enteritis,
before passing through the liver.38 Coughing, rhinorrhea, sore throat, malaise, and fever can develop
when the worms move through the lungs.110 Finally, the larvae reach the CNS after about 2 weeks and
eosinophilic meningitis and eosinophilic pleocytosis ensue.
The classical and most common clinical presentations of this disease are eosinophilic meningitis and
ocular angiostrongyliasis. However, a rare and extremely fatal encephalitic angiostrongyliasis was also
reported in some cases.111 The CNS damage is caused by direct mechanical and toxic injury from the
worm. The immunologic reactions of the host also play a role.112
The major pathological changes of human angiostrongyliasis occur in the brain.5 In autopsy studies, the external surface and spinal cord are generally normal and gross hemorrhage is not commonly seen. Infiltration of lymphocytes, plasma cells, and eosinophils is commonly reported in the
meninges and around intracerebral vessels.5 Cellular infiltration around living worms is not prominent, but dead worms are usually surrounded by a granuloma, increased number of eosinophils,
and sometimes Charcot–Leyden crystals.5 Although the clinical disease caused by Angiostrongylus
invasion into the CNS is commonly referred to as “eosinophilic meningitis,” the actual pathophysiology involves not just the meninges, or superficial lining of the brain, but also deeper brain tissue.
The physical lesions of tracks and microcavities caused by movement of the worms can be found in
the brain and even in the spinal cord. Initial invasion through the lining of the brain, the meninges,
may cause a typical inflammation of the meninges and a classic meningitis picture of headache, stiff
neck, and often fever accompanied by at least of one of the following symptoms: ataxia, face or limb
paralysis, visual disturbance, photophobia, hyperesthesia, or paresthesia.113 Aggregated data from
Thailand, Taiwan, mainland China, and the United States showed that 95% of patients suffered from
headache, 46% had mild neck stiffness, 44% suffered from persistent paresthesia, 38% had vomiting,
and 28% had nausea.19
The parasites subsequently invade deeper into the brain tissue, causing specific localizing neurologic
symptoms depending on where in the brain parenchyma they migrate. Continuous high intracranial pressure and corresponding damage to the brain and lung may precipitate unconsciousness, coma, and even
Biology of Foodborne Parasites
death in severe cases.114 Neurologic findings and symptoms wax and wane as initial damage is done by
the physical in-migration of the worms, and secondary damage is done by the inflammatory response to
the presence of dead and dying worms. This inflammation can lead in the short term to paralysis, bladder dysfunction, visual disturbance, and coma and in the long term to permanent nerve damage, mental
retardation, permanent brain damage, or death.19
The pathological findings in the CNS include the following: (1) meningitis with a predominance
of eosinophils and plasma cells; (2) tortuous tracks of various sizes in the brain and spinal cord surrounded by an inflammatory reaction and degenerating neurons; (3) granulomatous response to the
dead parasites; and (4) nonspecific vascular reactions including thrombosis, rupture of vessels, arteritis, and aneurysm formation.115 MRI may show prominence of Virchow–Robin spaces, periventricular hyperintense T2 signals, and enhancing subcortical lesions. Proton beam MR spectroscopy may
show decreased choline in the lesions.47 In the MRI findings with brain pathology, the hyperintense
signal intensity at the subcortical white matter in the T2-weighted and FLAIR imaging was probably
caused by nonspecific vascular or inflammatory reactions.116 The lesions represent focal parenchymal edema or demyelination caused by allergic or immunological reaction after invasion, death, and
protein dissolution of the nematode larvae. The disease appears to be the result of a vigorous host
response to larvae in the CNS. It can spontaneously resolve presumably because the larvae do not
survive in human hosts.
The clinical picture and symptoms of angiostrongyliasis in children differ greatly from those of adults.
Stiff neck and paresthesias are observed less frequently in children, but a high occurrence of nausea and
vomiting is found, with 82% of pediatric patients having nausea and vomiting. The incidence of fever
(up to 80%), somnolence (82%), constipation (76%), and abdominal pain (34.2%) is relatively higher in
children than among adults. It is possible that the increased severity in children is due to a higher worm
load relative to body size.19
A comprehensive analysis of human ocular angiostrongyliasis carried out in 2010 revealed that 1.2%
of all documented human cases were diagnosed with ocular angiostrongyliasis.37 Although the exact
mode of worm migration to eye is not known, the following route may be the most likely. After reaching
the brain tissue, larvae may migrate along the surface of the brain, particularly at the base of the brain,
where they may transverse the optic nerve, traveling between the nerve and sheath until they reach the
eye itself. The fifth-stage A. cantonensis larvae can be trapped between the optic nerve and nerve sheath
and in the periorbital tissue in rats.117 Another report of ocular angiostrongyliasis revealed the presence
of a nodule between the optic nerve and the sheath on orbital CT scan.118 Although it was not confirmed
by pathology, it is highly probable that the nodule was a dead larvae trapped between the optic nerve and
the sheath during the process of migration.
14.8 Treatment and Prevention
As relatively large number of rats and mollusks are highly susceptible to A. cantonensis worldwide, and
reports of other unnatural hosts are on a rise, it would be very difficult to eradicate this parasite from the
environment. However, it is possible to block the transmission of A. cantonensis to humans by educating
susceptible populations to avoid eating raw or undercooked intermediate and paratenic hosts or potentially contaminated vegetables.
In some of the endemic regions such as Thailand and China, the traditional custom of eating
snails prepared in various ways has been followed for many generations and became so popular that
it is extremely difficult to change food habits. Nonetheless, eating raw or undercooked snails with
seasonings such as pepper and pericarpium, which is very popular in Chinese restaurants, should
be strongly discouraged.5 Several outbreaks of human A. cantonensis infections in China have been
attributed to this method of preparing snails, and the simple approach to control the disease is to persuade people to abandon their habit of eating raw or undercooked intermediate and paratenic hosts
in endemic regions. Epidemiological surveys indicate that most cases of human angiostrongyliasis
would be avoided in this way. Some rare cases have been reported as caused by eating contaminated vegetables and vegetable juices. Thoroughly washing hands and utensils after preparing raw
snails or slugs or after gardening and intense washing of vegetables before cooking is recommended.
However, the difficulty for prevention is that most people have no or limited knowledge of the worm
and are totally unaware of the danger of consuming it.19 Therefore, increasing the public awareness
in endemic areas regarding the modes of transmission of A. cantonensis and its potential for damage
to the health of the general population is one of the most effective measures to control its spread.
Recommending adequate cooking before eating snails, slugs, small mollusks, and paratenic hosts of
A. cantonensis such as frogs, shrimps, land crabs, and monitor lizards and eradicating molluskan
hosts near houses and vegetable gardens are of prime importance in reducing the spread of this parasite in humans living in endemic areas.
Travelers heading to endemic regions must know the dangers of eating raw mollusks and raw vegetables with unknown sources and should avoid these foods. It is important that physicians in both nonendemic and endemic regions be aware of the existence of these worms, their symptoms and modes of
transmission, and methods to diagnose A. cantonensis infection in humans.
Most cases of human angiostrongyliasis exhibit two main forms of clinical presentations: eosinophilic meningitis and ocular angiostrongyliasis.19 The severity and clinical course of the disease depend
significantly on the ingested load of the third-stage larvae,36 making it difficult to design clinical trials
and to evaluate the effectiveness of treatments. Treatment is usually supportive with the use of analgesics for pain and corticosteroids to limit the inflammatory reaction. For eosinophilic meningitis,
effective supportive treatments are repeated lumbar puncture and analgesics.38,39 Typical conservative
medical management including analgesics and sedatives provides minimal relief to the headaches and
hyperesthesias. Removing CSF at 3- to 7-day intervals is the only proven method for significantly
reducing intracranial pressure and can be used for symptomatic treatment of headaches. This process
may be repeated until improvement is shown. Corticosteroid therapy has also been effective in human
One randomized, placebo-control study of a 2-week course of prednisolone (60 mg/day in three
divided doses) showed that the corticosteroids reduced the median length of headache from 13 to 5 days
and reduced the need for repeat lumbar puncture. Additionally, 9.1% of treatment patients compared to
45.5% of controls still had headache at 2 weeks.36
No antihelminthic drugs have been proven to be completely effective treatments, and there are some
concerns that antihelminthic could exacerbate neurological symptoms due to a systemic response to
dying worms. The effectiveness of any regimen may vary by endemic region. Albendazole and mebendazole have been used to treat this disease in attempts to more effectively relieve symptoms and reduce
their duration. The mean duration of headache was reduced significantly by using albendazole alone.120
The combination of corticosteroids and antihelminthics has been commonly used for treatment of human
angiostrongyliasis.121 Recent studies have shown that treatment using mebendazole or albendazole in
combination with prednisone or prednisolone can reduce the severity and duration of headaches but have
not been shown to improve long-term neurologic outcomes. In two small cases series (41 and 26 adults),
adult patients were given prednisolone 60 mg/day and an antihelminthic (mebendazole 10 mg/kg/day
or albendazole 15 mg/kg/day 2 weeks), resulting in resolution of headache in a median of 3 days in the
mebendazole case series and 4 days in the albendazole case series without serious side effects.121 Some
Chinese herbal medicines display efficacy for treating angiostrongyliasis in animal studies but have not
been used in humans.19
Several treatments, including corticosteroid, laser, and surgical removal of the parasite or a combination of therapies, have been used in treating ocular angiostrongyliasis. In a comprehensive review
of literature on ocular angiostrongyliasis,37 of 31 ocular angiostrongyliasis cases (where surgery was
performed to remove the worm), laser immobilization was carried out in 11 cases and retinal cryopexy in 2 cases prior to surgery. In some cases, intravenous methylprednisolone was administered to
lessen the intraocular inflammation and larvicidal drugs and steroid treatment were given in three cases.
Antihelminthic drugs, such as albendazole, are not usually recommended for the treatment because dead
worms may evoke severe inflammatory responses in the eye. However, regardless of treatment method,
the visual outcome of all 35 cases was not markedly improved from the presenting condition. Although
there is no evidence that surgical intervention improve the course of the disease, surgical removal is still
recommended to prevent further ocular damage.
Biology of Foodborne Parasites
14.9 Conclusion
Angiostrongyliasis is a foodborne disease and is transmitted to humans by consuming food contaminated with the larval stages of the parasite. However, most clinicians are not familiar with this disease
and little is known about the prevalence of A. cantonensis worldwide. Humans in endemic areas with
severe headache, stiff neck, nausea, vomiting, and paresthesia should be considered likely to be infected
with A. cantonensis, and parasitological and serological tests must be conducted to confirm or rule out
the tentative diagnosis.
Programs educating public health officials and physicians in endemic areas about A. cantonensis and its
hosts are practical and achievable interventions for the control of human infection. More studies should be
conducted on the biology of A. cantonensis and epidemiology and clinical characteristics of angiostrongyliasis, and more effective diagnostic methods and treatments for A. cantonensis should be developed.
A comprehensive analysis of the existing cases and outbreaks in the general population is also essential
to better understand its transmission. Educating the susceptible population regarding the mode of transmission and its clinical impact is the most important way to eradicate this foodborne disease. Completely
abstaining from eating the snail-associated delicacies, which is a key risk factor for A. cantonensis transmission, may not be possible. However, effective processing of these foods to kill the infective larvae
should be encouraged and practiced especially in highly endemic regions of this parasitic disease.
1. Chen, H.T., A new pulmonary nematode of rats, Pulmonema cantonensis ng, nsp from Canton, Ann
Parasitol, 13: 312–317, 1935. (in French.)
2. Nomura, S. and Lin, P.H., First case report of human infection with Haemostrongylus ratti Yokogawa,
Taiwan No Ikai, 3: 589–592, 1945.
3. Rosen, L., Laigret, J., and Boils, P.L., Observation on an outbreak of eosinophilic meningitis on Tahiti,
French Polynesia, Am J Hyg, 74: 26–42, 1961.
4. Li, H. et al., A severe eosinophilic meningoencephalitis caused by infection of Angiostrongylus cantonensis, Am J Trop Med Hyg, 79(4): 568–570, 2008.
5. Wang, Q.P. et al., Human angiostrongyliasis, Lancet Infect Dis, 8: 621–630, 2008.
6. New, D., Little, M.D., and Cross, J., Angiostrongylus cantonensis infection from eating raw snails,
N Engl J Med, 332: 1105–1106, 1995.
7. Bhaibulaya, M., A new species of Angiostrongylus in an Australian rat Rattus fuscipes, Parasitology, 58:
789–799, 1968.
8. Thiengo, S.C. et al., The giant African snail Achatina fulica as natural intermediate host of Angiostrongylus
cantonensis in Pernambuco, northeast Brazil, Acta Trop, 115: 194–199, 2010.
9. Mackerras, J.M. and Sandars, D.F., The life history of the rat lung-worm Angiostrongylus cantonensis
(Chen) (Nematoda: Metastrongylidae), Aust J Zool, 3: 1–21, 1955.
10. Alicata, J.E., The discovery of Angiostrongylus cantonensis as a cause of human eosinophilic meningitis,
Parasitol Today, 7: 151–153, 1991.
11. Jindrak, K., Angiostrongyliasis cantonensis (eosinophilic meningitis, Alicata’s disease), Contemp Neurol
Ser, 12: 133–164, 1975.
12. Lv, S. et al., Emerging angiostrongyliasis in mainland China, Emerg Infect Dis, 14: 161–164, 2008.
13. Barnes, R., Invertebrate Zoology, Orlando, FL: Dryden Press, 1987.
14. Brusca, R. and Brusca, G., Invertebrates, Sunderland, MA: Sinauer Associates, Inc., 2003.
15. Janovy, J. and Roberts, L., Foundations of Parasitology, 6th edn., Boston, MA: McGraw-Hill, 2000.
16. Anderson, D. et al., Eosinophilic meningoencephalitis due to Angiostrongylus cantonensis as the cause
of death in captive non-human primates, Am J Trop Med Hyg, 42: 70–74, 1990.
17. Prociv, P., Spratt, D., and Carlisle, M., Neuro-angiostrongyliasis: Unresolved issues, Int J Parasitol, 30,
1295–1303, 2000.
18. David, J.T. and Petri, W.A. Jr., Markell and Voge’s Medical Parasitology, St. Louis, MO: Elsevier, 2006.
19. Wang, Q.P. et al., Human Angiostrongylus cantonensis: An update, Eur J Clin Microbiol Infect Dis, 31,
389–395, 2012.
20. Alicata, J.E., Life cycle and biology, in: Alicata, J.E. and Jindrak, K. (eds.) Angiostrongyliasis in the
Pacific and Southeast Asia, Springfield, IL: C. C. Thomas, pp. 17–27, 1970.
21. Wu, G.H., Angiostrongylus cantonensis, in: Tang, J.Q. (ed.) Nature-Borne Diseases, Beijing, China:
Science Press, pp. 1182–1189, 2006. (in Chinese.)
22. Center for Disease Control and Prevention, Angiostrongylus cantonensis infection (online), Atlanta, GA:
CDC, Division of Parasitic Diseases, Parasitic Disease Information, 2013,
dpd/parasites/angiostrongylus/factsht_angiostrongylus.htm. Accessed March 11, 2013.
23. He, H. et al., Preliminary molecular characterization of the human pathogen Angiostrongylus cantonensis, BMC Mol Biol, 10, 97, 2009.
24. Fang, W. et al., ES proteins analysis of Angiostrongylus cantonensis: Products of the potential parasitism
genes? Parasitol Res, 106, 1027–1032, 2010.
25. Liu, Y.H. et al., Molecular cloning and characterization of cystatin, a cysteine protease inhibitor, from
Angiostrongylus cantonensis, Parasitol Res, 107, 915–922, 2010.
26. Hao, L. et al., Cloning, prokaryotic expression and immunoreactivity evaluation of Angiostrongylus cantonensis galectin, Nan Fang Yi Ke Da Xue Xue Bao, 27, 584–587, 2007.
27. Meng, J.X. et al., Identification and FQ-PCR investigation of GAMMA-BBH gene of Angiostrongylus
cantonensis, J Tropl Med, 7, 613–617, 2007.
28. Liu, L.H. et al., The mRNA level of the galectin-10 of Angiostrongylus cantonensis induced by reactive
oxygen stress, Parasitol Res, 112, 933–943, 2013.
29. Yang, X. et al., Molecular cloning, expression, and characterization of a putative activation-associated
secreted protein from Angiostrongylus cantonensis, Parasitol Res, 112, 781–788, 2013.
30. Monte, T.C. et al., Phylogenetic relationship of the Brazilian isolates of the rat lungworm Angiostrongylus
cantonensis (Nematoda: Metastrongylidae) employing mitochondrial COI gene sequence data, Parasitol
Res, 5, 248, 2012.
31. Thaenkham, U. et al., Population structure of Angiostrongylus cantonensis (Nematoda: Metastrongylidae)
in Thailand based on PCR-RAPD markers, Southeast Asian J Trop Med Public Health, 43(3), 567–573,
32. Liu, Q. et al., Molecular characterization and immunolocalization of a protein disulfide isomerase from
Angiostrongylus cantonensis, Parasitol Res, 110, 2501–2507, 2012.
33. Cheng, M. et al., Cloning and characterization of a novel cathepsin B-like cysteine proteinase from
Angiostrongylus cantonensis, Parasitol Res, 110, 2413–2422, 2012.
34. Chang, S.H., Tang, P., and Wang, L.C., A transcriptomic study on the pepsin-activated infective larvae of
Angiostrongylus cantonensis, Mol Biochem Parasitol, 179, 47–50, 2011.
35. Chen, M.X. et al., Angiostrongylus cantonensis: Identification and characterization of microRNAs in
male and female adults, Exp Parasitol, 128, 116–120, 2011.
36. Tsai, H.C. et al., Eosinophilic meningitis caused by Angiostrongylus cantonensis: Report of 17 cases, Am
J Med, 111, 109–114, 2001.
37. Zongli, D. et al., Human ocular angiostrongyliasis: A literature review, Tropical Doctor, 41, 76–78, 2011.
38. Yii, C.Y., Clinical observations on eosinophilic meningitis and meningoencephalitis caused by
Angiostrongylus cantonensis in Taiwan, Am J Trop Med Hyg, 25, 233–249, 1976.
39. Punyagupta, S., Juttijudata, P., and Bunnag, T., Eosinophilic meningitis in Thailand. Clinical studies of
484 typical cases probably caused by Angiostrongylus cantonensis, Am J Trop Med Hyg, 24, 921–931,
40. Slom, T.J. et al., An outbreak of eosinophilic meningitis caused by Angiostrongylus cantonensis in travelers returning from the Caribbean, N Engl J Med, 346, 668–675, 2002.
41. Kuberski, T. and Wallace, G.D., Clinical manifestations of eosinophilic meningitis due to Angiostrongylus
cantonensis, Neurology, 29, 1566–1570, 1979.
42. Jin, E. et al., MRI findings of eosinophilic myelomeningoencephalitis due to Angiostrongylus cantonensis, Clin Radiol, 60, 242–250, 2005.
43. Ogawa, K. et al., A case of eosinophilic meningoencephalitis caused by Angiostrongylus cantonensis
with unique brain MRI findings, Rinsho Shinkeigaku 38, 22–26, 1998. (in Japanese.)
44. Hasbun, R. et al., Computed tomography of the head before lumbar puncture in adults with suspected
meningitis, N Engl J Med, 345, 1727–1733, 2001.
45. Chau, T.T. et al., Headache and confusion: The dangers of a raw snail supper, Lancet, 361, 1866, 2003.
46. Lo, R.V. and Gluckman, S.J., Eosinophilic meningitis, Am J Med, 114, 217–223, 2003.
Biology of Foodborne Parasites
47. Kanpittaya, J. et al., MR findings of eosinophilic meningoencephalitis attributed to Angiostrongylus cantonensis, AJNR Am J Neuroradiol, 21, 1090–1094, 2000.
48. Tsai, H.C. et al., Eosinophilic meningitis caused by Angiostrongylus cantonensis associated with eating
raw snails: Correlation of brain magnetic resonance imaging scans with clinical findings, Am J Trop Med
Hyg, 68, 281–285, 2003.
49. Hsu, W.Y. et al., Eosinophilic meningitis caused by Angiostrongylus cantonensis, Pediatr Infect Dis J, 9,
443–445, 1990.
50. Shih, S.L. et al., Angiostrongylus cantonensis infection in infants and young children, Pediatr Infect Dis
J, 11, 1064–1066, 1992.
51. Eamsobhana, P., Yoolek, A., and Kreethapon, N., Blinded multi-laboratory evaluation of an in-house dotblot ELISA kit for diagnosis of human parastrongyliasis, Southeast Asian J Trop Med Public Health, 34,
1–6, 2003.
52. Eamsobhana, P. et al., Detection of circulating antigens of Parastrongylus cantonensis in human sera by
dot-blot ELISA and sandwich ELISA using monoclonal antibody, Southeast Asian J Trop Med Public
Health, 28, 624–628, 1997.
53. Morassutti, A.L. et al., The 31-kDa antigen of Angiostrongylus cantonensis comprises distinct antigenic
glycoproteins, Vector Borne Zoonotic Dis, 12, 961–968, 2012.
54. Zhang, X. et al., Analysis of larval excretory-secretory antigen and its immunodiagnosis of
Angiostrongyliasis cantonensis infection, Nan Fang Yi Ke Da Xue Xue Bao, 32, 477–481, 2012.
55. Eamsobhana, P. and Yong, H.S., Immunological diagnosis of human angiostrongyliasis due to
Angiostrongylus cantonensis (Nematoda: Angiostrongylidae), Int J Infect Dis, 13, 425–430, 2009.
56. Chen, M.X. et al., Monoclonal antibodies against excretory/secretory antigens of Angiostrongylus cantonensis, Hybridoma, 29, 447–452. 2010.
57. Huang, D.N. et al., Detection of circulating antigen of Angiostrongylus cantonensis by 12D5 and 21B7
monoclonal antibodies, Chinese J Epidemiol, 31, 79–82, 2010.
58. Chen, M.X. et al., Development of a double antibody sandwich ELISA assay for the diagnosis of angiostrongyliasis, J Parasitol, 97, 721–724, 2011.
59. Chen, M.X. et al., Seroprevalence of Angiostrongylus cantonensis infection in humans in China,
J Parasitol, 97, 144–145, 2011.
60. Chye, S.M. et al., Immuno-PCR for detection of antigen to Angiostrongylus cantonensis circulating fifthstage worms, Clin Chem, 50, 51–57, 2004.
61. Maleewong, W. et al., Immunoblot evaluation of the specificity of the 29-kDa antigen from young adult
female worms Angiostrongylus cantonensis for immunodiagnosis of human angiostrongyliasis, Asian
Pac J Allergy Immunol, 19, 267–273, 2001.
62. Nuamtanong, S., The evaluation of the 29 and 31 kDa antigens in female Angiostrongylus cantonensis
for serodiagnosis of human angiostrongyliasis, Southeast Asian J Trop Med Public Health, 27, 291–296,
63. Bessarab, I.N. and Joshua, G.W., Stage-specific gene expression in Angiostrongylus cantonensis:
Characterisation and expression of an adult-specific gene, Mol Biochem Parasitol, 88, 73–84, 1997.
64. Intapan, P.M. et al., Evaluation of human IgG subclass antibodies in the serodiagnosis of angiostrongyliasis, Parasitol Res, 89, 425–429, 2003.
65. Tavares, R.G. et al., Molecular techniques for the study and diagnosis of parasite infection, J Venom Anim
Toxins Incl Trop Dis, 17, 239–224, 2011.
66. Silva, A.C.A., Graeff-Teixeira, C., and Zaha, A., Diagnosis of abdominal angiostrongyliasis by PCR
from sera of patients, Rev Inst Med Trop Sao Paulo, 45, 295–297, 2003.
67. Eamsobhana, P. et al., Molecular differentiation and phylogenetic relationships of three Angiostrongylus
species and Angiostrongylus cantonensis geographical isolates based on a 66-kDa protein gene of A. cantonensis (Nematoda: Angiostrongylidae), Exp Parasitol, 126, 564–569, 2010.
68. Eamsobhana, P. et al., Molecular diagnosis of eosinophilic meningitis due to Angiostrongylus cantonensis (Nematoda: Metastrongyloidea) by polymerase chain reaction-DNA sequencing of cerebrospinal
fluids of patients), Mem Inst Oswaldo Cruz, Rio de Janeiro, 108, 116–118, 2013.
69. Wei, F.R. et al., Multiplex PCR assay for the detection of Angiostrongylus cantonensis larvae in Pomacea
canaliculata, Zhongguo Ji Sheng Chong Xue Yu Ji Sheng Chong Bing Za Zhi, 28, 355–358, 2010.
70. Liu, C.Y. et al., Specific detection of Angiostrongylus cantonensis in the snail Achatina fulica using a
loop-mediated isothermal amplification (LAMP) assay, Mol Cell Probes, 25, 164–167, 2011.
71. Qvarnstrom, Y. et al., Improved molecular detection of Angiostrongylus cantonensis in mollusks and
other environmental samples with a species-specific internal transcribed spacer 1-based TaqMan assay,
Appl Environ Microbiol, 76, 5287–5289, 2010.
72. Jaryi, S.I. et al., Quantitative PCR estimates Angiostrongylus cantonensis (rat lungworm) infection levels
in semi-slugs (Parmarion martensi), Mol Biochem Parasitol, 185, 174–176, 2012.
73. Chen, J.X. et al., A protein microarray for the rapid screening of patients suspected of infection with various food-borne helminthiases, PLoS Negl Trop Dis, 6, e1899, 2012.
74. Cross, J.H. and Chen, E.R., Angiostrongyliasis, in: Murrell, K.D. and Fried, B. (eds.) Food-Borne
Parasitic Zoonoses, New York: Springer, pp. 263–290, 2007.
75. Schmutzhard, E., Boongird, P., and Vejjajiva, A., Eosinophilic meningitis and radiculomyelitis in
Thailand, caused by CNS invasion of Gnathostoma spinigerum and Angiostrongylus cantonensis,
J Neurol Neurosurg Psychiatry, 51, 80–87, 1988.
76. Chen, X.G., Li, H., and Lun, Z.R., Angiostrongyliasis, Mainland China, Emerg Infect Dis, 11, 1645–
1647, 2005.
77. Zheng, R.Y. et al., Probing and demonstrating etiological factors for outbreak of Angiostrongyliasis cantonensis in Wenzhou, Sh J Prev Med, 13, 105–107, 2001.
78. Lin, W. and Wang, X.T., Epidemiology of Angiostrongylus cantonensis in mainland, Chin J Zoonoses,
20, 1004–1007, 2004. (in Chinese.)
79. Yang, F.Z. et al., Survey on the outbreak of human angiostrongyliasis caused by eating snails, Strait
J Prev Med, 10, 44–45, 2004. (in Chinese.)
80. Wu, C.H. and Yan, X.H., Outbreak of human angiostrongyliasis in Fuzhou, Chin J Zoonoses, 20, 454,
81. Lin, J.X. et al., Epidemiological study on group infection of Angiostrongylus cantonensis in Changle
City, Zhongguo Ji Sheng Chong Xue Yu Ji Sheng Chong Bing Za Zhi, 21, 110–112, 2003. (in Chinese.)
82. Chen, W.L. et al., Eosinophilic meningitis: 31 cases report, Chin J Mis Diagn, 6, 4668–4669, 2006.
83. Han, J.H. et al., Eosinophilic meningitis: 28 cases report, J Pathol Biol, 1, 2–3, 2006.
84. Chen, F. et al., Investigation on outbreak of angiostrongyliasis cantonensis due to consumption of snail
food in Dali City, Zhongguo Xue Xi Chong Bing Fang Zhi Za Zhi, 23, 687–690, 2011.
85. Deng, Z.H. et al., An outbreak of angiostrongyliasis in Guanging, People’s Republic of China: Migrants
vulnerable to an emerging disease, Southeast Asian J Trop Med Public Health, 42, 1047–1053, 2011.
86. Pascual, G.J.E., Aguilar, P.P.H., and Galvez, O.M.D., Finding of Angiostrongylus cantonensis in the cerebrospinal fluid of a boy with eosinophilic meningoencephalitis, Rev Cubana Med Trop, 33, 92–95, 1981.
87. Vazquez, J.J. et al., Angiostrongyliasis in a European patient: A rare cause of gangrenous ischemic
enterocolitis, Gastroenterology, 105, 1544–1549, 1993.
88. Lindo, J.F. et al., Fatal autochthonous eosinophilic meningitis in a Jamaican child caused by
Angiostrongylus cantonensis, Am J Trop Med Hyg, 70, 425–428, 2004.
89. Brown, F.M., Mohareb, E.W., Yousif, F., Sultan, Y., and Girgis, N.I., Angiostrongylus eosinophilic meningitis in Egypt, Lancet, 348, 964–965, 1996.
90. Lv, S. et al., Invasive snails and an emerging infectious disease: Results from the first national survey on
Angiostrongylus cantonensis in China, PLoS Negl Trop Dis, 3, e368, 2008.
91. Kliks, M.M. and Palumbo, N.E., Eosinophilic meningitis beyond the Pacific basin: The global dispersal
of a peridomestic zoonosis caused by Angiostrongylus cantonensis, the nematode lungworm of rats, Soc
Sci Med, 34, 199–212, 1992.
92. Tsai, T.H. et al., An outbreak of meningitis caused by Angiostrongylus cantonensis in Kaohsiung,
J Microbiol Immunol Infect, 34, 50–56, 2001.
93. Toma, H. et al., Ocular angiostrongyliasis without meningitis symptoms in Okinawa, Japan, J Parasitol,
88, 211–213, 2002.
94. Wang, Q.P., Chen, X.G., and Lun, Z.R., Invasive freshwater snail, China, Emerg Infect Dis, 13, 1119–
1120, 2007.
95. Zhang, R.L. et al., Enzootic angiostrongyliasis in Shenzhen, China, Emerg Infect Dis, 14, 1995–1996,
96. Liang, H.K., Shen, H.X., and Xu, B.K., Investigation on the definite, intermediate and paratenic hosts of
Angiostrongylus cantonensis in Guangzhou, Chin J Epidemiol, 5, 245–248, 1984.
97. Yang, X. et al., Enzootic angiostrongyliasis in Guangzhou, China, 2008–2010, Am J Trop Med Hyg, 86,
846–849, 2012.
Biology of Foodborne Parasites
98. Nowak, R. and Paradiso, J., Walker’s Mammals of the World, 4th edn., Baltimore, MD: The Johns
Hopkins University Press, 1983.
99. Silver, J., The introduction and spread of house rats in the United States, J Mammalogy, 8, 58–60, 1927.
100. Lunn, J.A. et al., Twenty two cases of canine neural angiostronglyosis in eastern Australia (2002–2005)
and a review of the literature, Parasite Vectors, 5, 70, 2012.
101. Tsai, H.C. et al., Outbreak of eosinophilic meningitis associated with drinking raw vegetable juice in
southern Taiwan, Am J Trop Med Hyg, 71, 222–226, 2004.
102. Ash, L.R., The occurrence of Angiostrongylus cantonensis in frogs of New Caledonia with observations
on paratenic hosts of metastrongyles, J Parasitol, 54, 432–436, 1968.
103. Radomyos, P. et al., Occurrence of the infective stage of Angiostrongylus cantonensis in the yellow tree
monitor (Varanus bengalensis) in five provinces of Thailand, Southeast Asian J Trop Med Public Health,
25, 498–500, 1994.
104. Panackel, C. et al., Eosinophilic meningitis due to Angiostrongylus cantonensis, Indian J Med Microbiol,
24, 220–221, 2006.
105. Hidelaratchi, M.D., Riffsy, M.T., and Wijesekera, J.C., A case of eosinophilic meningitis following monitor lizard meat consumption, exacerbated by anthelminthics, Ceylon Med J, 50, 84–86, 2005.
106. Gardiner, C.H. et al., Eosinophilic meningoencephalitis due to Angiostrongylus cantonensis as the cause
of death in captive non-human primates, Am J Trop Med Hyg, 42, 70–74, 1990.
107. Gemma, M. et al., Tawny frogmouths and brushtail possums as sentinels for Angiostrongylus cantonensis, the rat lungworm, Vet Parasitol, 192, 158–165, 2013.
108. He, Z.Y. et al., Survey on the outbreak of human angiostrongyliasis in Beijing, Chin J Public Health, 23,
1241–1242, 2007. (in Chinese.)
109. Xue, D.Y. et al., Epidemiological investigation on an outbreak of Angiostrongyliasis cantonensis in
Wenzhou, Zhongguo Ji Sheng Chong Xue Yu Ji Sheng Chong Bing Za Zhi, 18, 176–178, 2000. (in Chinese.)
110. Cross, J.H., Clinical manifestations and laboratory diagnosis of eosinophilic meningitis syndrome associated with angiostrongyliasis, Southeast Asian J Trop Med Public Health, 9, 161–170, 1978.
111. Sawanyawisuth, K. et al., Clinical factors predictive of encephalitis caused by Angiostrongylus cantonensis, Am J Trop Med Hyg, 81, 698–701, 2009.
112. Pipitgool, V. et al., Angiostrongylus infections in rats and snails in northeast Thailand, Southeast Asian
J Trop Med Public Health, 28, 190–193, 1997.
113. Tsai, H.C. et al., Brain magnetic resonance imaging abnormalities in eosinophilic meningitis caused by
Angiostrongylus cantonensis infection, Vector Borne Zoonotic Dis, 12, 2, 2012.
114. Wang, J. et al., An outbreak of Angiostrongyliasis cantonensis in Beijing, J Parasitol, 96, 377–381, 2010.
115. Nye, S.W. et al., Lesions of the brain in eosinophilic meningitis, Arch Pathol Lab Med, 89, 9–19, 1970.
116. Jin, E.H. et al., Magnetic resonance imaging of eosinophilic meningoencephalitis caused by
Angiostrongylus cantonensis following eating freshwater snails, Chin Med J (Engl), 121, 67–72, 2008.
117. Xiang, D.M. et al., A case of ocular angiostrongyliasis in a child, Chin J Strab Pediat Ophthalmol, 9, 91,
2001. (in Chinese)
118. Qi, H., Diao, Z., and Yin, C., A case of optic nerve compression caused by Angiostrongylus cantonensis,
Am J Trop Med Hyg, 81, 4, 2009.
119. Chotmongkol, V., Sawanyawisuth, K., and Thavornpitak, Y., Corticosteroid treatment of eosinophilic
meningitis, Clin Infect Dis, 31, 660–662, 2000.
120. Jitpimolmard, S. et al., Albendazole therapy for eosinophilic meningitis caused by Angiostrongylus cantonensis, Parasitol Res, 100, 1293–1296, 2007.
121. Chotmongkol, V. et al., Treatment of eosinophilic meningitis with a combination of prednisolone and
mebendazole, Am J Trop Med Hyg, 74, 1122–1124, 2006.
Simonetta Mattiucci, Michela Paoletti, Paolo Cipriani,
Stephen C. Webb, and Giuseppe Nascetti
15.1 Introduction................................................................................................................................... 255
15.2 Morphology and Classification..................................................................................................... 256
15.3 Molecular Methods for Characterization of Anisakis spp............................................................ 257
15.3.1 Multilocus Allozyme Electrophoresis.............................................................................. 257
15.3.2 DNA Sequencing.............................................................................................................. 257
15.3.3 PCR–RFLP Analysis........................................................................................................ 257
15.4 Biology of Anisakis....................................................................................................................... 257
15.5 Pathogenesis and Clinical Features.............................................................................................. 266
15.5.1 Gastric Anisakiasis........................................................................................................... 266
15.5.2 Intestinal Anisakiasis....................................................................................................... 266
15.5.3 Gastroallergic Anisakiasis............................................................................................... 267
15.5.4 Anisakis Allergy............................................................................................................... 267
15.6 Diagnosis....................................................................................................................................... 268
15.6.1 Molecular Diagnosis......................................................................................................... 268
15.6.2 Serodiagnosis.................................................................................................................... 269
15.7 Treatment and Prevention............................................................................................................. 269
References............................................................................................................................................... 269
15.1 Introduction
Nematodes belonging to the genus Anisakis are heteroxenous parasites that use marine mammals (mainly
cetaceans) as definitive hosts and crustaceans (krill), fish, and squid as intermediate/paratenic hosts in
their life cycles. These nematodes are of medical and economic concerns due to their public health
importance and effects on the marketability of fish products. Larval Anisakis spp. are the main causative
agents of a fish-borne parasitic zoonosis named “anisakiasis,” via accidental ingestion of raw or undercooked fish or squid infected by those larvae. In humans, Anisakis larvae do not develop to the adult
stage in the alimentary tract but may penetrate the gastrointestinal tract, often with severe pathological
consequences, such as the formation of eosinophilic granulomas. In addition, evidence is accumulating
that these parasites can induce strong allergic reactions.
The prevalence of human infection is highest in countries where consumption of raw fish is ­widespread.
Human cases are increasingly reported in Japan, many European countries (the United Kingdom, France,
Spain, and Italy), and elsewhere. To assess the impact of these parasites on human health, epidemiological studies on anisakid infections in fish and squid are needed. Much has been done to assess the
prevalence of Anisakis infection in edible fish, and risk maps depicting geographic areas, transmission
season, host size, and other important parameters have been compiled. Public health agencies such as the
European Food Safety Authority (EFSA)1 have also taken an interest in this fish-borne zoonosis.
Biology of Foodborne Parasites
This chapter briefly reviews the systematics of Anisakis species, with biological and ecological data
including life cycle, definitive/intermediate/paratenic hosts, geographical distribution, and their occurrence in edible parts of the fish. In addition, human disease due to Anisakis spp., its clinical forms, and
diagnosis are also described. The chapter concludes with a review of research needs to improve our
knowledge on this human health threat.
15.2 Morphology and Classification
Morphological characters of taxonomic significance in nematodes belonging to the genus Anisakis are
very few and often only applicable to adult parasites (such as the number and distribution of male caudal papillae, position of the vulva, and length of the spicules),2–5 making impossible their identification
based only on morphology, especially in the case of “sibling species” (morphologically identical but
reproductively isolated taxa). Indeed, speciation events in anisakid nematodes occur generally without
morphological differentiation, and the adaptation to a stable environment has led to convergence of morphological features. The paucity of diagnostic characters is even more marked in larvae. Thus, Anisakis
larvae can be identified morphologically only to genus level, mainly on the basis of the shape and length
of the glandular part of the esophagus (ventriculus) and the absence of the intestinal cecum and ventricular appendix. Two morphotypes of Anisakis spp., Type I and Type II, may be discerned by the presence/
absence of a caudal spine (mucron) and the long/short ventriculus.6 Epizootiological and epidemiological
studies, to be meaningful, require correct identification of the etiological agent. However, in Anisakis
studies, morphology is not particularly useful.
Molecular approaches allow accurate identification of Anisakis to the species level. Thus, inferences
about their systematics and evolution of anisakid nematodes using these techniques are of obvious
implications for epidemiology. Multilocus allozyme electrophoresis (MAE) has revealed the existence
of extensive genetic heterogeneity within anisakid morphospecies, such as Anisakis simplex s.l.7 The
biological species concept (BSC)8 was well supported by the application of allozyme markers for several
Anisakis species. Indeed, the diversity of species belonging to Anisakis has increased after the detection
of several sibling species, leading to the discovery and description of several new species. Reproductive
isolation and the absence of gene flow between sympatric and allopatric sibling species have been demonstrated by allozymes, thus establishing their specific status.2–5,7,9–11
DNA-based approaches for species identification, such as PCR restriction fragment length polymorphism (PCR–RFLP) and direct sequencing of ITS ribosomal DNA (rDNA)12 or mitochondrial
genes,13 have confirmed the previous taxonomic assessment of Anisakis spp. Currently, allozymes,14
PCR–RFLP of rDNA,12,15 and DNA sequence analyses of nuclear (ITS region of the rDNA)15,16 and
mitochondrial (mtDNA cytochrome oxidase 2 [cox2] and rrnS) genes 4–5,13,15,16 have demonstrated that
the genus Anisakis comprises of at least nine distinct species. They include the three species in the
A. simplex s.l. complex, that is, A. simplex (Rudolphi, 1809) (sensu stricto), A. pegreffii (CampanaRouget and Biocca, 1955) (= A. simplex A7), A. berlandi (Mattiucci et al., 2014) (= A. simplex C9;
the two closely related taxa, A. ziphidarum 2 and A. nascettii 4), A. typica (Diesing, 1860), and other
three closely related species, A. physeteris (Baylis, 1920), A. brevispiculata (Dollfus, 1966), and
A. paggiae.3
The existence of two major clades (Clade I and Clade II) has been demonstrated by phylogenetic
inferences.4,5,13,15 Clade I comprises of one subclade formed by A. simplex (sensu stricto), A. pegreffii,
and A. berlandi and a second one containing A. ziphidarum and A. nascettii. In contrast, three species
belong to Clade II, including A. physeteris, A. brevispiculata, and A. paggiae. A. typica forms a distinct
phylogenetic lineage divergent from other species.5,15
So far, Anisakis spp. included in the Clade I exhibit morphotype I at the larval stage, whereas Anisakis
spp. in Clade II exhibit morphotype II. Thus, at present, five species of Anisakis share Type I morphology, while three other species show larval morphotype II. Therefore, the larval stages of Anisakis
spp. cannot be identified to species by means of morphological features without the use of molecular
In addition to the nine species of Anisakis characterized so far, recent studies17,18 have detected
genetically the existence of one additional taxon, recovered as a larval stage in a nonmigratory
fish species from Balinese, Javanese, and Malaysian waters of the Pacific Ocean. Preliminary data
suggest that the taxon (Anisakis sp. 1) may be a sibling species of A. typica occurring in central
Pacific waters.18 A further gene pool, referred to as Anisakis sp. 2, has been detected genetically
by allozyme and mtDNA cox2 sequence analysis based on Type II larvae from swordfish in the
equatorial area.19,20
Despite the limited morphological characters available in adult Anisakis spp., morphological and morphometric studies have found some diagnostic features useful for species determination. Morphological
differences have been detected in A. paggiae with respect to the closely related taxa A. ­brevispiculata
and A. physeteris.10 Similarly, some morphological features have distinguished A. nascettii from
A. ­ziphidarum14 and allowed differentiation of A. pegreffii, A. simplex (s.s.), and A. berlandi.5,21
15.3 Molecular Methods for Characterization of Anisakis spp.
As emphasized earlier, the limited value of morphological analysis makes essential the use of molecular
methods in the identification of species of the genus Anisakis. Some of the currently used molecular/
genetic methods are described here briefly.
15.3.1 Multilocus Allozyme Electrophoresis
MAE (19–24 enzyme loci) has been extensively used in identifying the large number of Anisakis populations sampled from many geographic regions from the Boreal and Austral hemispheres, detecting
“sibling species,” discovering new species, and addressing questions concerning the population genetics,
evolutionary biology, and the relationship between genetic variability and habitat disturbance.18 The currently used diagnostic allozymes to identify all the species of Anisakis are described by Mattiucci et al.4
15.3.2 DNA Sequencing
DNA sequencing of some genes is useful for the identification of the different species in siblings and
morphospecies of Anisakis. Sequences of nuclear and mitochondria genes are available for almost all the
nine species recognized within the genus Anisakis.
For the analysis of mitochondrial DNA, two regions have been sequenced in all Anisakis taxa: cox24,5,13
and rrnS (the small subunit of the rDNA).5,16 These mitochondrial markers can distinguish all taxa genetically characterized so far. The mtDNA cox2 region shows a high degree of polymorphism at the intraspecific level; this finding supports the use of this gene in further studies of the population genetics and
phylogeography.22,23 For the analysis of nuclear genes, the rDNA region spanning the final part of the 18S
subunit, the first internal transcribed spacer (ITS-1), the 5.8S subunit, the second internal transcribed
spacer (ITS-2), and the beginning of the 28S subunit have been sequenced for all known Anisakis spp.5,15
15.3.3 PCR–RFLP Analysis
PCR–RFLP analysis12 of the ITS-1 and ITS-215,16 has been extensively used for the identification of
Anisakis species.
15.4 Biology of Anisakis
Anisakis species have an indirect, complex life cycle, which involves various marine organisms at
­different levels of a trophic web in the marine ecosystem. The adults live in the stomachs of marine
mammals, mainly cetaceans. The life cycle begins when females release eggs, which are passed through
Biology of Foodborne Parasites
the feces of their definitive host, into the sea. According to some experimental studies, the eggs embryonate, producing third-stage larvae.26–28 They are ingested by crustaceans such as copepods and euphausiids (krill), in which they grow in the hemocoel. Fish or squid (Cephalopoda, Decapodiformes) become
infected after eating an infected crustacean; the third-stage larvae bore into the digestive tract wall of
the fish and pass into the visceral body cavity of the fish to undergo induced encapsulation.29 The life
cycle is completed after the intermediate/paratenic host (fish, squid, or, directly, crustaceans) is eaten by
a definitive host; inside the stomach or the intestine of its mammalian host, Anisakis spp. develop into
sexually mature female or male nematodes.
Because Anisakis larvae do not undergo any development or molt inside the fish or squid, these hosts
should be regarded as paratenic. Small fishes and squid are frequently eaten by larger fish, thus generating an additional passage in a new paratenic host. This is important from an epidemiological point of
view because the repeated transmission of Anisakis larvae in the prey–predator system allows extensive
bioaccumulation of the parasites in larger fish. Several pelagic and demersal fish species show increased
prevalence and abundance of Anisakis larvae with age and size.29–31 In infected fish, most of the larvae
occur in the visceral body cavity, typically encapsulated outside the organs. However, some larvae may
migrate from the visceral cavity to the musculature of the fish, mostly in the belly flap, but also in the
dorsal musculature. It has been demonstrated that in some fish, the migration occurs “intra vitam,” rather
than “postmortem” of the fish species.32 The occurrence of larvae in fish fillets is a human health hazard.
It has been suggested that different species of Anisakis larvae could have a different capacity to migrate
and infect fish fillets.33,34
A. simplex (sensu stricto) is widespread between 35 °N and the Arctic Circle; it is present in
both the western and eastern Atlantic and Pacific Oceans9,16,35–42 (Figure 15.1). The southern limit
of this species seems to be in the northeast Atlantic Ocean in waters around the Gibraltar area.
A. ­simplex (s.s.) is also occasionally identified at the larval stage in western Mediterranean waters
olar C
Arctic P
A. pegreffii
A. simplex (s.s.)
A. berlandi (= A. simplex C)
A. typica
A. physeteris
A. brevispiculata
A. paggiae
A. nascettii
FIGURE 15.1 Distribution of Anisakis spp. in the world.
A. ziphidarum
due to the migration of pelagic fish species into the Alboran Sea from the Atlantic.18,19,30 Adult
worms parasitize several species of cetacean hosts (Table 15.2), while several fish and squid species
have been found harboring larvae of this species throughout its geographical range. A sympatric
area between A. simplex (s.s.) and A. pegreffii has been identified along the Spanish and Portuguese
Atlantic coasts,9,19,30,36,43–46 in the Alboran Sea,19,30 and in Japanese Sea waters.37,41,42 A. simplex (s.s.)
also occurs in sympatry with A. berlandi in the eastern Pacific Ocean, where it has been identified
in definitive and intermediate/paratenic hosts.2,5,9,35 Several oceanic dolphins in the Delphinidae,
Arctic dolphins in the Monodontidae, and porpoises in the Phocoenidae (Table 15.2) are hosts of
A. simplex (s.s.).9,18,47
A. pegreffii is the most important anisakid nematode in several pelagic and demersal fish from
Mediterranean waters.2,18,48–52 Adults and larvae are also widely distributed in the Austral Region
between 35 °N and 55 °S.5,9 In Atlantic waters, the northern limit of its geographical range seems to be
the Iberian coast and Portuguese coast.9,19,30,36,44–46 It has been detected at the larval stage in some fish
from Japanese37,38,40–42 and Chinese waters.53 Some oceanic dolphins in the Delphinidae (Table 15.2) are
hosts of A. pegreffii.9,18,47
A. berlandi (A. simplex C9) exhibits a discontinuous range, including the Canadian and Chilean
Pacific coasts, New Zealand waters, and the South African Atlantic coast.9,16 This species has been
identified at the adult stage syntopically with A. pegreffii in cetaceans and as a larva in some fish species (Table 15.1). It has been occasionally identified in Mirounga leonina from the sub-Antarctic area18
and in M. ­angustirostris from the northeast Pacific Ocean.16 Some oceanic dolphins in the Delphinidae
(Table 15.2) are hosts of A. berlandi.5,9,18
The range of A. typica extends from 30 °S to 35 °N in warmer temperate and tropical waters.11,50 In
these areas, it is found as an adult in dolphins and as larvae in several fish species (Tables 15.1 and 15.2).
A. typica has also been identified in cetaceans and fish from the eastern Mediterranean Sea (off Cyprus).
Its presence in these waters could be the result of “Lessepsian migration” (through the Suez Canal)30 of
its intermediate/paratenic hosts from the Indian Ocean. It was identified in flatfish captured in central
Portuguese waters of the Northeast Atlantic Ocean45 and rarely in some fish caught along the North
African coast (Tunisia and Libya) of the Mediterranean Sea.48 A. typica larvae were recognized also in
fish from Chinese waters.53
A. ziphidarum2 was first identified genetically in beaked whales from the South Atlantic Ocean (off
the South African coast) and in the Mediterranean Sea. A. ziphidarum has also been found in Central
Atlantic waters up to the Caribbean Sea50,54 and in South Pacific waters (New Zealand coast).47 Thus,
it appears to have a wide geographical range related to that of its definitive hosts. Little information is
available concerning its infection in fish and/or squid; some larvae were recognized in some fish species
from central Atlantic waters.30,44,46 However, it seems that this species may involve other intermediate
hosts, such as squid, rather than fish in its life cycle, as these animals represent the main food source of
beaked whales.18
A. nascettii4 has been detected at the adult stage in beaked whales from New Zealand waters and
from the South African coast. It was also identified as an L4 stage in ziphiids from the central Atlantic
Ocean.55 This species has been genetically identified, at the larval stage, in the squid Moroteuthis ingens
in the Tasman Sea. This appears to support the hypothesis that this species involves squid rather than
finfish in its life cycle.4
A. physeteris was first genetically characterized in its main definitive host, the sperm whale,
Physeter macrocephalus, from Mediterranean waters,56 and genetically identified adults have also
been recorded in the central Atlantic Ocean.15,18 Type II larvae of A. physeteris have been genetically identified in the swordfish, Xiphias gladius, from Mediterranean and Atlantic waters.20
A. ­brevispiculata was initially characterized genetically using allozymes,10 the mtDNA cox2 gene,13
and by ITS rDNA sequence analysis12,16 based on parasite material from pygmy sperm whales, Kogia
breviceps, from South African and Northeast Atlantic waters (Iberian coast) (Table 15.2). Anisakis
larvae of Type II corresponding to A. brevispiculata were recognized by allozyme markers as a
rare parasite in the fish Merluccius merluccius30 and heavily infecting the swordfish X. gladius in
­t ropical–equatorial Atlantic waters.20
Biology of Foodborne Parasites
TABLE 15.1
Intermediate/Paratenic Hosts of Anisakis spp. Detected Using Molecular Techniques
(s.s.) pegreffii berlandi typica ziphidarum nascettii physeteris brevispiculata paggiae
Sepia officinalis
Illex coindetii
Illex illecebrosus
M. ingens
Todarodes pacificus
Todaropsis eblanae
Anoplopoma fimbria
Belone belone
Beryx splendens
Arnoglossus laterna
Brama brama
Trachurus capensis
Trachurus picturatus
Trachurus trachurus
Deania profundorum
Caesio cuning
Citharus linguatula
(Continued )
TABLE 15.1 (Continued )
Intermediate/Paratenic Hosts of Anisakis spp. Detected Using Molecular Techniques
(s.s.) pegreffii berlandi typica ziphidarum nascettii physeteris brevispiculata paggiae
Clupea harengus
Sardinops sagax
Conger conger
nitidus nitidus
Boreogadus saida
Gadus morhua
Trisopterus luscus
Thyrsites atun
Gerres oblongus
Etmopterus princeps
Etmopterus pusillus
Etmopterus spinax
Lampris guttatus
Lophius piscatorius
Lophius vomerinus
Molva dypterygia
Brosme brosme
(Continued )
Biology of Foodborne Parasites
TABLE 15.1 (Continued )
Intermediate/Paratenic Hosts of Anisakis spp. Detected Using Molecular Techniques
(s.s.) pegreffii berlandi typica ziphidarum nascettii physeteris brevispiculata paggiae
Pinjalo lewisi
Pinjalo pinjalo
Merluccius capensis
Merluccius hubbsi
Muraena helena
Nemipterus virgatus
Genypterus capensis
pretiosus japonicus
Phycis phycis
Phycis blennoides
Parapercis colias
Platichthys flesus
(Continued )
TABLE 15.1 (Continued )
Intermediate/Paratenic Hosts of Anisakis spp. Detected Using Molecular Techniques
(s.s.) pegreffii berlandi typica ziphidarum nascettii physeteris brevispiculata paggiae
Petromyzon marinus
Oncorhynchus keta
Salmo salar
Scomberesox saurus
Auxis rochei rochei
Auxis thazard
Euthynnus affinis
Sarda orientalis
Scomber japonicus
Scomber scombrus
Thunnus albacares
Thunnus thynnus
Scorpaena scrofa
(Continued )
Biology of Foodborne Parasites
TABLE 15.1 (Continued )
Intermediate/Paratenic Hosts of Anisakis spp. Detected Using Molecular Techniques
(s.s.) pegreffii berlandi typica ziphidarum nascettii physeteris brevispiculata paggiae
Echiichthys vipera
Trachinus draco
Aphanopus carbo
Eutrigla gurnardus
X. gladius
Note: Data are from Refs. [2,3,7,9–11,14,18,20,30,33,36–38,44–46,48,50–51,53,56,96,107–116]. Hosts are listed by
alphabetical order of the family.
A. paggiae was first genetically characterized and described morphologically as an adult parasite of
the pygmy sperm whale and the dwarf sperm whale (Table 15.2) off both Florida and the South African
Atlantic coasts.3 It was identified in kogiids from the Caribbean Sea.15 In recent years, this parasite was
identified also in Kogia sima from Japanese Sea waters.57
The phylogenetic relationships proposed elsewhere18,47 for the species of Anisakis “mirror,” the phylogeny proposed so far for their cetacean definitive hosts.18,58–61
TABLE 15.2
Definitive Hosts of Anisakis spp. Detected by Molecular Techniques
(s.s.) pegreffii berlandi typica ziphidarum nascettii physeteris brevispiculata paggiae
Delphinus capensis
Delphinus delphis
Globicephala melas
Lagenodelphis hosei
Orcinus orca
Sotalia fluviatilis
Sotalia guianensis
Stenella clymene
Stenella attenuata
Stenella longirostris
Steno bredanensis
Tursiops truncatus
K. breviceps
K. sima
Caperea marginata
Phocoena phocoena
P. macrocephalus
(Continued )
Biology of Foodborne Parasites
TABLE 15.2 (Continued )
Definitive Hosts of Anisakis spp. Detected by Molecular Techniques
(s.s.) pegreffii berlandi typica ziphidarum nascettii physeteris brevispiculata paggiae
Mesoplodon grayi
Mesoplodon mirus
Ziphius cavirostris
M. leonina
Note: Data are from Refs. [2–3,7,9–11,14,15,18,30,41,50,54,56,112,116,117]. Hosts listed by alphabetical order of the family.
15.5 Pathogenesis and Clinical Features
The L3 stage of Anisakis spp. infecting the flesh of sea fish or squid can be ingested alive by humans,
causing the zoonotic disease anisakiasis. First reported in the Netherlands, anisakiasis has gained an
increasing health and economic relevance in particular in countries such as Japan, where the consumption of raw fish and squid is frequent, although human cases are increasingly reported in many European
countries (Spain, Italy, the United Kingdom, and France). A number of fish dishes are considered to be
high risk for the contraction of human anisakiasis. They include Scandinavian gravlax, Dutch salted and
marinated herrings, Japanese sushi and sashimi, Spanish boquerones and anchovies, and Italian marinated anchovies.62 Despite these reports, it is likely that anisakiasis is still an underestimated zoonosis.
Among the nine species of Anisakis described earlier, only two, that is, A. simplex (s.s.) and A. pegreffii,
are reported so far as causative agents of human anisakiasis.38,63–67
Depending on the location of the ingested Anisakis larva in humans, the disease is subdivided into gastric anisakiasis (GA) and intestinal anisakiasis (IA). It has been classified as “acute” or “moderate” by the
symptoms related to the two clinical forms and “not invasive” or “invasive” depending on whether the larva
remains in the gastric and/or intestinal lumen or invades the submucosal layer of the gastric or intestinal wall.
The acute form is mostly gastric, and it is characterized by nausea, vomiting, and epigastric pain,
which appear approximately 1–6 h after ingestion of infected fish. In IA, acute signs could appear about
7 days after infection with abdominal pain, nausea, vomiting, fever, diarrhea, and fecal occult blood.
Several rarely occurring extragastrointestinal localizations have been also documented (oropharyngeal,
abdominal cavity, mesenteries, and omentum).
15.5.1 Gastric Anisakiasis
Epigastric pain, nausea, vomiting, abdominal fullness or distension, and anorexia are the most frequent
symptoms in acute GA. Endoscopic examination may reveal edematous hypertrophic gastric folds, an
increase in gastric secretion and peristalsis, and mucosal lesions, including edema, redness, coagulation,
hemorrhage, and ulceration. GA described recently from Italy67 was characterized by epigastric pain 2 h
after eating raw seafood. Endoscopy revealed that Anisakis larvae were mainly located in the lumen of
the stomach and not invading the submucosal layer of the gastric wall.
15.5.2 Intestinal Anisakiasis
The “mild form” of IA is characterized by “tumorlike” eosinophilic granulomas in the intestinal walls,
whereas the “fulminant form” has symptoms of acute ileus, acute appendicitis, and acute abdominal
and regional ileitis. Cases of IA reported from Italy64,66 were characterized by a clinical picture of the
“fulminant form” with acute abdominal pain and acute appendicitis. Radiography or ultrasonography
images show increased thickness of the intestinal wall, a marked dilatation of the intestine, the so-called
“keyboard sign” could be present; in addition, ascites pooling between the dilated intestine could appear,
in both the “mild” and “fulminant” forms of IA.68
15.5.3 Gastroallergic Anisakiasis
This is an acute allergic reaction associated with the gastric presence of Anisakis larvae, when they
attempt to invade the submucosal layer of the gastric wall.69 Gastroallergic anisakiasis (GAA) consists
of acute IgE-mediated generalized reaction with urticaria, angioedema, and anaphylaxis. In this type
of anisakiasis, allergic reactions may begin 2–3 h to 2–3 days after the ingestion of infected fish.67,69
Recently, A. pegreffii was implicated (by molecular methods) in two GAA cases, characterized by
­urticaria and edema of the oral mucosa in Italian patients after they had consumed “marinated anchovies.”67 Endoscope examination showed the larvae invading the submucosal layer of the gastric wall. In
addition, serum samples from these patients showed IgE reactivity in western blotting (WB) analysis
against Ani s 1 antigen of A. pegreffii.
15.5.4 Anisakis Allergy
Anisakis allergy was described first in Japan68 and has been recently reviewed.69,70 Allergic reactions
associated with high levels of IgE have been reported after the ingestion of infected fish fillets, even
after the removal of larvae.62 In Spain, since 1995, more than 150 cases of allergy due to Anisakis have
been reported.62,71 Additionally, cases of occupational Anisakis allergy including the detection of high
IgE level were described in fishermen or associated with exposure (by either contact or inhalation) to
fish meal.72,73 Clinical symptoms of allergic anisakiasis range from urticaria to anaphylactic shock. The
diagnosis of allergic anisakiasis is in general associated with high levels of IgE and the recognition of
specific antigens/allergens in WB.
To date, 12 Anisakis (Ani s) allergens have been characterized, numbered from Ani s 1 to Ani s 12
(Table 15.3). They include both somatic (S) and excretory/secretory (E/S) antigens, some of which remain
ill defined (Table 15.3). The major allergens of Anisakis, which are recognized by IgE and IgG in serum
samples of patients, are Ani s 1 (24 kDa) and Ani s 7 (139 kDa); they are located in E/S glands.62 Ani s 1
TABLE 15.3
Allergens of Anisakis (Ani s)
Ani s 1
Ani s 1 isoform
Ani s 2
Ani s 3
Ani s 4
Ani s 5
Ani s 6
Ani s 7
Ani s 8
Ani s 9
Ani s 10
Ani s 11
Ani s 11-li
Ani s 12
MW (kDa)
Location of the Products
Major Allergen
S (?)
S (?)
S (?)
Note: E/S, excretory/secretory products; S, somatic; ?, unknown location in the larva or molecular weight.
Biology of Foodborne Parasites
was recognized by IgE in sera of GAA patients.67,74 Ani s 1 has also been detected in a high percentage
of Anisakis-sensitized patients from Morocco.72 Ani s 1 seems to be a heat-stable allergen74; therefore,
allergic reactions could occur after consumption of not only undercooked fresh fish but also cooked or
frozen infected fish. There is also an isoform (at 21 kDa) of Ani s 1.75 It has been suggested that the “mild”
form of GA and allergic conditions in Spain are related to this isoform of Ani s 1.74,75 Ani s 2 and Ani s
3 allergens are somatic (muscular) proteins in the Anisakis larva; they are paramyosin and tropomyosin
and are considered “panallergens.” A phylogenetic comparison of tropomyosin amino acid sequences
demonstrated that nematode tropomyosins of Anisakis and Ascaris are closely related to those of insects,
crustaceans, and mites.70 This indicates a possible immunological cross-reactivity. Indeed, the Anisakis
muscle proteins, paramyosin and tropomyosin, are thought to be responsible for the cross-reactivity
between Anisakis and other invertebrates and for the IgE hypersensitivity detection in blood sera often
reported in allergic patients.76,77
Ani s 7, a glycoprotein, is also considered to be a major allergen, being recognized in up to 100% of
sera samples in patients with Anisakis allergy.78 The amino acid sequence similarity of Ani s 1, Ani s 7,
and Ani s 12 in A. pegreffii and A. simplex (s.s.) has recently been described.79 Other minor allergens
are represented by Ani s 4, a heat-stable protein, which is recognized by 27%–30% of patients,80 and
Ani s 5, Ani s 8, and Ani s 9, which are all heat-stable E/S proteins less frequently recognized in patient
sera.81–83 These allergens can be considered food allergens. Additional allergens have been identified, namely, Ani s 10–12; however, little is known about their function and location in the Anisakis
larvae. Furthermore, a hemoglobin from A. pegreffii was recently identified as being responsible for
high immunoactivity in hypersensitive patients; it has a phylogenetic similarity to other invertebrate
Finally, healthy individuals can have high levels of anti-Anisakis IgE in their serum without the development of allergic symptoms. On the other hand, individuals with low levels of specific IgE antibodies
may show clinical manifestations of anisakiasis. In one study, a patient with minimal symptoms of
allergy was seen by endoscopy to have an intense parasite burden (200 Anisakis larvae) in the stomach84
yet the specific humoral response to Anisakis was weak. This is congruent with a previous experimental
model85 in which a high parasite load could lead to a poor IgE response, suggesting a possible immunomodulation role played by Anisakis larvae.
15.6 Diagnosis
15.6.1 Molecular Diagnosis
Despite the fact that human infection is high in countries where eating raw fish is widespread, the
molecular identification of human cases remains scarce. Most reports come from European countries
(especially from the southern Mediterranean countries) where allergic symptoms and hypersensitivity
associated with the parasite are frequently reported.
As mentioned earlier, morphology is insufficient to identify larval Anisakis to species. Furthermore,
when larvae infect humans, specimens collected are often spoiled or fragmented, making identification impossible even to genus level. Molecular methodologies, in contrast, do not have these limitations. Anisakis larvae removed by endoscopy from GA cases were identified by PCR amplification of
the entire ITS and subsequent RFLP analysis.63,65,86,87 Six cases of GA and two cases of GAA identified by endoscopy in eight Italian patients were diagnosed as belonging to the same species, that is,
A. ­pegreffii, by PCR amplification and sequencing of the ITS region and the mtDNA cox2 gene by
Mattiucci et al.67 Furthermore, molecular methods have also allowed the identification of A. pegreffii in
a paraffin-­embedded granuloma associated with IA.66
Molecular identification of specimens from human anisakiasis cases has so far detected only the sibling species A. pegreffii and A. simplex (s.s.). No data are available concerning the occurrence of the third
species of the A. simplex complex, A. berlandi (A. simplex C). We conclude from human cases so far
diagnosed by using molecular markers that A. pegreffii is able to cause gastric and intestinal anisakiasis
and GAA in humans.67
15.6.2 Serodiagnosis
Currently, serodiagnostic tests for Anisakis include the use of ImmunoCAP systems, immunoblotting
(WB), ELISA, and skin prick tests (SPTs).62 All of these methods use partially purified or crude extracts
of Anisakis larvae. This makes these methodologies of poor specificity, due to their cross-reactivities with
antigens from other parasites and allergens. Therefore, it is possible that IgE detection by ImmunoCAP
assay can overestimate the number of human cases sensitized to Anisakis allergens. Some researchers
have used IgE and IgG detection in immunoblotting (WB) to differentiate, for instance, anisakiasis and
Anisakis allergy from asymptomatic Anisakis IgE-sensitized patients.71,88,89 Several purified Anisakis
allergens, when used in combination, have proven to be useful in diagnosis by WB.90
15.7 Treatment and Prevention
Endoscopic removal of larvae in GA and the surgical treatment of granulomas in IA resolve the disease.
The efficacy of antihelminthics is not supported by large surveys, although recent studies have shown
significant in vitro actions of albendazole against Anisakis larvae.90 This seems to support the possible
use of this drug in treating clinical manifestations of human anisakiasis, at least when the Anisakis larvae are still in the stomach, a few hours after the ingestion of the infected fish.
The consumption of fish infected by larvae of the genus Anisakis poses a biological hazard that can be
mitigated by control measures under the supervision of health authorities and by proper storage and processing methods that allow the inactivation of the larvae. Immediate evisceration of fish may reduce the
zoonotic potential of the parasite by preventing migration of worms into the flesh of host fish. However,
this practice is not useful when the migration of the larvae occurs while the fish is alive.
In many countries where fish fillets are heavily infected, fish are examined during their processing.91
A usual procedure for the larval detection in fish fillets is candling on the light table; fillets are sliced
and observed by transmitted light. However, this method is inefficient, and it has been estimated that it is
only able to detect ~33% of heavily infected fish fillets.91 In recent years, the preferred detection methods
of anisakids in fish products (both fresh and processed) are based on pepsin digestion92 and compression
of fillets and observation with ultraviolet light.93–95 Other methods include real-time PCR analysis of ITS
and mtDNA cox1 genes.96,97 However, these methods are not yet validated and have been used for the
detection of only two species of Anisakis (i.e., A. simplex s.s. and A. pegreffii). A quantitative sandwich
ELISA assay for the detection of Anisakis protein in seafood has been evaluated.98
For the prevention of human anisakiasis, cooking the fish at 70°C quickly kills larvae. However, some
allergens in fish fillets (e.g., Ani s 1, Ani s 4, Ani s 9) appear to be thermostable, and allergens released
from the larvae into the surrounding tissue may retain their allergenicity even after death of the larvae
by heat treatments.99 Regarding deep freezing, Adams et al.100 reported low survival of larvae per fillet (0%–3%) after 6 h at −40°C, but up to 30% survived after 48 h at −20°C. Similarly, Wharton and
Aalders101 demonstrated that larvae can survive at temperatures down to −10°C. Deep freezing (−20°C
for 24 h) and cooking for sufficiently periods are the most effective methods. Cold smoking and marinating do not kill the larvae, unless high concentrations of food-grade acetic acid are used.102 Dry salting
can kill the parasite, provided that the salt is evenly distributed in all parts of the muscle and is used
at correct concentrations (>20 baumè, see Ref. [103]). Recently, Brutti et al.104 demonstrated complete
inactivation of A. simplex larvae in raw fish using high-hydrostatic-pressure treatments, while the effect
of microwave treatments has been reported by Adams et al.,105 Tejada et al.,106 and Vidaček et al.99
1. EFSA, Panel on Biological hazards (BIOHAZ). Scientific opinion on risk assessment of parasites in
fishery products, EFSA Journal, 8(4), 1541, 2010.
2. Paggi, L. et al., A new species of Anisakis Dujardin, 1845 (Nematoda: Anisakidae) from beaked whales
(Ziphiidae): Allozyme and morphological evidence, Syst. Parasitol., 40, 161, 1998.
3. Mattiucci, S. et al., Evidence for a new species of Anisakis Dujardin, 1845: Morphological description
and genetic relationships between congeners (Nematoda: Anisakidae), Sys. Parasitol., 61, 157, 2005.
Biology of Foodborne Parasites
4. Mattiucci, S. et al., Anisakis nascettii n. sp. (Nematoda: Anisakidae) from beaked whales of the southern
hemisphere: Morphological description, genetic relationships between congeners and ecological data,
Syst. Parasitol., 74, 199, 2009.
5. Mattiucci, S. et al., Genetic and morphological approaches distinguishing the three sibling species of the
Anisakis simplex species complex, with a species designation as Anisakis berlandi n. sp. for A. simplex
sp. C (Nematoda: Anisakidae), J. Parasitol., 100(2), 199, 2014. doi:
6. Berland, B., Nematodes from some Norwegian marine fishes, Sarsia, 2, 1, 1961.
7. Nascetti, G. et al., Electrophoretic studies on the Anisakis simplex complex (Ascaridida: Anisakidae)
from the Mediterranean and North-East Atlantic, Int. J. Parasitol., 16, 633, 1986.
8. Mayr, E., Animal Species and Evolution. Cambridge, MA: Belknap Press, Harvard University Press,
p. 797, 1963.
9. Mattiucci, S. et al., Genetic and ecological data on the Anisakis simplex complex, with evidence for a new
species (Nematoda, Ascaridoidea, Anisakidae), J. Parasitol., 86, 401, 1997.
10. Mattiucci, S. et al., Genetic divergence and reproductive isolation between Anisakis brevispiculata and
Anisakis physeteris (Nematoda: Anisakidae), Int. J. Parasitol., 31, 9, 2001.
11. Mattiucci, S. et al., Genetic markers in the study of A. typica (Diesing, 1860): Larval identification
and genetic relationships with other species of Anisakis Dujardin, 1845 (Nematoda: Anisakidae), Syst.
Parasitol., 51, 159, 2002.
12. D’Amelio, S. et al., Genetic markers in ribosomal DNA for the identification of members of the genus
Anisakis (Nematoda: Ascaridoidea) defined by polymerase chain reaction-based restriction fragment
length polymorp