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CRYPTOSPORIDIUM AND
CRYPTOSPORIDIOSIS Second Edition
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CRYPTOSPORIDIUM AND
CRYPTOSPORIDIOSIS Second Edition
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CRYPTOSPORIDIUM AND
CRYPTOSPORIDIOSIS Second Edition
Edited by
Ronald Fayer Lihua Xiao
Boca Raton London New York
CRC Press is an imprint of the Taylor & Francis Group, an informa business
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CRC Press Taylor & Francis Group 6000 Broken Sound Parkway NW, Suite 300 Boca Raton, FL 33487-2742 © 2008 by Taylor & Francis Group, LLC CRC Press is an imprint of Taylor & Francis Group, an Informa business No claim to original U.S. Government works Printed in the United States of America on acid-free paper 10 9 8 7 6 5 4 3 2 1 International Standard Book Number-13: 978-1-4200-5226-8 (Hardcover) This book contains information obtained from authentic and highly regarded sources. Reprinted material is quoted with permission, and sources are indicated. A wide variety of references are listed. Reasonable efforts have been made to publish reliable data and information, but the author and the publisher cannot assume responsibility for the validity of all materials or for the consequences of their use. No part of this book may be reprinted, reproduced, transmitted, or utilized in any form by any electronic, mechanical, or other means, now known or hereafter invented, including photocopying, microfilming, and recording, or in any information storage or retrieval system, without written permission from the publishers. For permission to photocopy or use material electronically from this work, please access www.copyright.com (http:// www.copyright.com/) or contact the Copyright Clearance Center, Inc. (CCC) 222 Rosewood Drive, Danvers, MA 01923, 978-750-8400. CCC is a not-for-profit organization that provides licenses and registration for a variety of users. For organizations that have been granted a photocopy license by the CCC, a separate system of payment has been arranged. Trademark Notice: Product or corporate names may be trademarks or registered trademarks, and are used only for identification and explanation without intent to infringe. Library of Congress Cataloging-in-Publication Data Cryptosporidium and cryptosporidiosis / [edited by] Ronald Fayer, Lihua Xiao. -- 2nd ed. p. ; cm. “A CRC title.” Includes bibliographical references and index. ISBN 978-1-4200-5226-8 (hardcover : alk. paper) 1. Cryptosporidiosis. 2. Cryptosporidium. I. Fayer, R. II. Xiao, Lihua, 1962- III. Title. [DNLM: 1. Cryptosporidiosis. 2. Cryptosporidium. WC 730 C956 2007] RC136.5.C79 2007 616.9’36--dc22 Visit the Taylor & Francis Web site at http://www.taylorandfrancis.com and the CRC Press Web site at http://www.crcpress.com
2007021158
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Ernest Edward Tyzzer (Courtesy of the Francis A. Countway Library of Medicine, Boston, Massachusetts)
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Contents
1
General Biology ....................................................................................................1 Ronald Fayer
2
Genomics .............................................................................................................43 Jessica C. Kissinger
3
Biochemistry .......................................................................................................57 Guan Zhu
4
Epidemiology ......................................................................................................79 Gordon Nichols
5
Molecular Epidemiology ................................................................................. 119 Lihua Xiao and Una M. Ryan
6
Diagnostics ........................................................................................................ 173 Huw Smith
7
Immune Responses .......................................................................................... 209 Vincent McDonald
8
Clinical Disease and Pathology...................................................................... 235 Cirle Alcantara Warren and Richard L. Guerrant
9
Prophylaxis and Chemotherapy...................................................................... 255 Heather D. Stockdale, Jennifer A. Spencer, and Byron L. Blagburn
10 Foodborne Transmission.................................................................................. 289 Ynes R. Ortega and Vitaliano A. Cama
11
Waterborne: Drinking Water ........................................................................... 305 Jennifer L. Clancy and Thomas M. Hargy
12 Waterborne: Recreational Water ..................................................................... 335 Michael J. Beach
13 Waste Management........................................................................................... 371 Dwight D. Bowman
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14 Fish, Amphibians, and Reptiles ..................................................................... 387 Thaddeus K. Graczyk
15 Birds ................................................................................................................... 395 Una M. Ryan and Lihua Xiao
16 Zoo and Wild Mammals .................................................................................. 419 Olga Matos
17 Companion Animals ........................................................................................ 437 Mónica Santín and James M. Trout
18 Livestock............................................................................................................ 451 Mónica Santín and James M. Trout
19 Animal Models ................................................................................................. 485 Saul Tzipori and Giovanni Widmer
20 In Vitro Cultivation .......................................................................................... 499 Michael J. Arrowood Index ........................................................................................................................... 527
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Preface
In the century following E.E. Tyzzer’s pioneering description of Cryptosporidium in mice, the genus Cryptosporidium has been overlooked, rediscovered, and now found to consist of numerous species and genotypes adapted to parasitic life in virtually all classes of vertebrates. In the past decade our knowledge has expanded from microscopic observations of infection and environmental contamination to the knowledge acquired from widespread application of molecular techniques to taxonomy and epidemiology, the sequencing of the genome of two major species, and greater understanding of the biochemistry and phylogeny of members of this fascinating genus. This second edition of Cryptosporidium and Cryptosporidiosis has been greatly revised and expanded in response to the volume and scope of new information on these parasites of human and veterinary importance, and the need to provide a comprehensive and up-to-date treatment consolidating the thousands of scientific reports. Interest in Cryptosporidium has spread from its academic base among biologists and parasitologists to veterinarians, physicians, epidemiologists, pharmacologists, public health specialists, drinking water and waste water managers, swimming pool managers, farmers, backpackers, and the public in general. Concern for prevention and treatment extends from underdeveloped communities to highly industrialized societies, for immunocompromised persons as well as healthy populations, for persons of all ages from infants to the elderly, and for persons caring for animals from companion animals and livestock to captive exotic species. Chapter 1 discusses general biological issues. It traces the history of discovery of the genus and species, updates the taxonomy, describes the life cycle stages and their morphology, addresses host specificity, and summarizes factors that reduce oocyst transmission. Chapter 2 introduces molecular biology to the study of Cryptosporidium through description of data types, properties of the genome, genetic regulation, and comparative genomics. Chapter 3 updates the biochemistry of this genus. It describes energy and carbohydrate metabolism as well as nucleotide, fatty acid, polyamine, amino acid, and DNA and RNA metabolism. It describes structural proteins, membrane proteins and transporters, and delves into potential drug targets. Chapter 4 provides epidemiologists and persons interested in transmission dynamics with detailed descriptions of outbreaks and the methods used to trace the sources. Chapter 5 is devoted to molecular epidemiology. It describes the molecular tools, the population genetics of Cryptosporidium species, the epidemiology of animal and human infections, and tracking sources in water. Chapter 6 provides laboratory technicians and diagnosticians detailed descriptions of the vast array of tests used for oocyst recovery, concentration, and purification as well as microscopic methods for staining and observing oocysts. It summarizes immunological and molecular methods for detecting infection and, when possible, identifying species. Chapter 7 describes host immune responses. The innate immunity of epithelial cells and natural killer cells, T-cell-mediated immunity, parasite-specific priming of cells, the roles of cytokines, protection from parasite specific antibodies, and vaccination against infection are discussed. Chapter 8 provides medical workers with the description of clinical disease and pathology in humans, including infections in immunocompetent and immunocompromised persons, and age-related infections. It discusses organ sites, histopathology, pathophysiology, and human volunteer studies. Chapter 9 details the vast array of compounds tested with emphasis on those found most effective for prophylaxis and treatment of cryptosporidiosis in humans and animals. Chapter 10 reviews foodborne transmission, the outbreaks, methods of detection in various foods, sources of contamination, decontamination, and HACCP and other regulations. Chapter 11 presents the issues concerned with Cryptosporidium oocysts in drinking water. These include outbreaks, methods for detection and prevention, and government regulations related to the drinking water industry. Chapter 12 discusses factors associated with transmission of Cryptosporidium in various types of recreational waters. It describes the outbreaks and means to reduce or prevent future outbreaks. Chapter 13 deals with waste management, specifically: treatment of wastewater effluent, sludge treatment, and treatment of manure from cattle and swine.
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Chapter 14 reviews infections in fish, amphibians, and reptiles, and discusses treatment, prevention, and control. Chapter 15 addresses cryptosporidiosis in birds: the disease, immunity, prophylaxis, and treatment; cultivation of the parasite; and the major species and genotypes infecting birds. Chapter 16 covers the range of wild animals infected with Cryptosporidium from rodents, lagomorphs, insectivores, omnivores, ruminants, carnivores, bats, and marsupials to nonhuman primates. Chapter 17 discusses cryptosporidiosis of major companion animal species: cats, dogs, and horses. Chapter 18 discusses cryptosporidiosis of major livestock species: cattle, sheep, goats, pigs, and other species. Chapter 19 provides new information on animal models best suited for parasite propagation: the rodent, pig, monkey, and gerbil models best suited for research; and models for testing parasite–host range. And finally, Chapter 20 provides descriptions of in vitro methods of studying Cryptosporidium, the cells and media found most useful, and the techniques for producing and storing purified parasites. The editors acknowledge with gratitude the chapter authors for their contributions to this book. Ronald Fayer Lihua Xiao
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Editors
Ronald Fayer received his B.S. degree from the University of Alaska–Fairbanks and his M.S. and Ph.D. degrees under the direction of Professor Datus Hammond at Utah State University. Dr. Fayer began his professional career as a zoologist in the Beltsville Parasitology Laboratory of the U.S. Department of Agriculture’s Agricultural Research Service developing in vitro cultivation methods for the protozoan parasites Eimeria, Toxoplasma, Besnoitia, Isospora, Hepatozoon, and Sarcocystis. Following up on the discovery of the coccidian life cycle of Sarcocystis in cell culture, studies were devoted to defining the pathological effects in hosts including immunopathological and metabolic perturbations resulting in heart lesions, abortions, retarded growth, and mortality. Preceding the advent of AIDS, his research emphasis shifted to Cryptosporidium and has encompassed molecularly based surveys of prevalence in livestock; effects of disinfectants, heat and cold on oocysts; immune responses in mouse models and calves; passive immunotherapy with colostral antibody; drug treatments; mechanical transport hosts; identification of new species using molecular tools; and use of molluscan shellfish as bio-indicators of fecal pollution of surface waters. Dr. Fayer has served in a variety of administrative capacities in the Agricultural Research Service including research leader, laboratory chief, national program leader for parasitology and toxicology, director of the Animal Parasitology Institute, and assistant area director for the Northeastern states. He has served on the editorial boards of five scientific journals and as president of the Helminthological Society of Washington, the American Association of Veterinary Parasitologists, and the American Society of Parasitologists. He was the recipient of a senior Fulbright Fellowship. He has published over 325 papers in scientific journals and five books. For his contributions to research he has received the H.B. Ward Medal from the American Society of Parasitologists, the National Distinguished Scientist of the Year Award from the Agricultural Research Service, the Superior Service Award and the Plow Award from the U.S. Department of Agriculture, and the Presidential Rank Award for Distinguished Senior Professional in the career civil service. Lihua Xiao obtained his veterinary education in China. After receiving his M.S. in veterinary parasitology and teaching for 2 years at the Northeast Agricultural University in Harbin, China, he obtained his Ph.D. in veterinary parasitology under Professor Harold Gibbs at the University of Maine and received postdoctoral training with Professor Rupert Herd at the Ohio State University College of Veterinary Medicine. In 1993, he moved to the Centers for Disease Control and Prevention (CDC), first as a guest researcher, then as a senior staff fellow. He is currently a senior scientist in the Division of Parasitic Diseases, National Center for Zoonotic, Vector-Borne, and Enteric Diseases, CDC, Atlanta, Georgia. Dr. Xiao’s earlier research interests were mostly on the epidemiology, pathogenesis, and control of gastrointestinal nematodes of farm animals. Prior to the massive cryptosporidiosis outbreak in Milwaukee, his research shifted to epidemiology and biology of cryptosporidiosis and giardiasis of farm animals. More recently, he has focused on the taxonomy, molecular epidemiology, and environmental biology of Cryptosporidium, Giardia, microsporidia, and other enteric protists in humans and animals, while working simultaneously on the immunopathogenesis and vaccine development of malaria and immunopathogenesis of HIV. Dr. Xiao has published over 200 scientific papers, invited reviews, and book chapters, over half of which are on Cryptosporidium and cryptosporidiosis. He has received the James H. Nakano citation from the National Center for Infectious Diseases, CDC, and the Outstanding Overseas Young Scientist Award from the National Science Foundation of China. He also holds several adjunct faculty positions at universities in the United States and China.
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Contributors
Michael J. Arrowood Centers for Disease Control and Prevention, Atlanta, Georgia Michael J. Beach Centers for Disease Control and Prevention, Atlanta, Georgia Byron L. Blagburn
College of Veterinary Medicine, Auburn University, Auburn, Alabama
Dwight D. Bowman Department of Microbiology and Immunology, College of Veterinary Medicine, Cornell University, Ithaca, New York Vitaliano A. Cama Bloomberg School of Public Health, Johns Hopkins University, Baltimore, Maryland Jennifer L. Clancy Clancy Environmental Consultants, Inc., St. Albans, Vermont Ronald Fayer United States Department of Agriculture, Agricultural Research Service, Animal and Natural Resources Institute, Environmental Microbial Safety Laboratory, Beltsville, Maryland Thaddeus K. Graczyk Bloomberg School of Public Health, Johns Hopkins University, Baltimore, Maryland Richard L. Guerrant Center for Global Health, Division of Infectious Diseases and International Health, University of Virginia School of Medicine, Charlottesville, Virginia Thomas M. Hargy Clancy Environmental Consultants, Inc., St. Albans, Vermont Jessica C. Kissinger Center for Tropical and Emerging Global Diseases, and Department of Genetics, University of Georgia, Athens, Georgia Olga Matos Unit of Opportunistic Protozoa/HIV and Other Protozoa, Institute of Hygiene and Tropical Medicine, New University of Lisbon, Portugal Vincent McDonald Centre for Gastroenterology, Barts and the London School of Medicine, Queen Mary College, London, United Kingdom Gordon Nichols Centre for Infections, Health Protection Agency, London, United Kingdom Ynes R. Ortega Center for Food Safety and Department of Food Science and Technology, University of Georgia, Griffin, Georgia Una M. Ryan School of Veterinary and Biomedical Sciences, Murdoch University, Perth, Australia Mónica Santín United States Department of Agriculture, Agricultural Research Service, Animal and Natural Resources Institute, Environmental Microbial Safety Laboratory, Beltsville, Maryland Huw Smith Scottish Parasite Diagnostic Laboratory, Stobhill Hospital, Glasgow, Scotland
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Jennifer A. Spencer
College of Veterinary Medicine, Auburn University, Auburn, Alabama
Heather D. Stockdale College of Veterinary Medicine, Auburn University, Auburn, Alabama James M. Trout United States Department of Agriculture, Agricultural Research Service, Animal and Natural Resources Institute, Environmental Microbial Safety Laboratory, Beltsville, Maryland Saul Tzipori Cummings School of Veterinary Medicine, Tufts University, North Grafton, Massachusetts Cirle Alcantara Warren Center for Global Health, Division of Infectious Diseases and International Health, University of Virginia School of Medicine, Charlottesville, Virginia Giovanni Widmer Cummings School of Veterinary Medicine, Tufts University, North Grafton, Massachusetts Lihua Xiao Division of Parasitic Diseases, Center for Disease Control and Prevention, Atlanta, Georgia Guan Zhu Department of Veterinary Pathobiology, College of Veterinary Medicine Biomedical Sciences, Texas A&M University, College Station, Texas
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1 General Biology
Ronald Fayer
CONTENTS I. II. III.
Introduction ...................................................................................................................................... 1 History .............................................................................................................................................. 2 Taxonomy ......................................................................................................................................... 4 A. Requirements for Species Identification............................................................................. 6 1. Valid and Nonvalid Species ................................................................................. 7 IV. Host Specificity .............................................................................................................................. 10 V. Life Cycle....................................................................................................................................... 11 A. The Environmental Stage and Ultimate Source of Infection........................................... 12 B. Excystation ........................................................................................................................ 13 C. Cell Invasion and Internalization ...................................................................................... 13 D. Asexual Multiplication (Merogony) ................................................................................. 16 E. Sexual Reproduction (Gamogony).................................................................................... 16 F. Sporogony ......................................................................................................................... 18 G. Prepatent Patent Times...................................................................................................... 19 H. Extracellular Stages........................................................................................................... 20 VI. Structure ......................................................................................................................................... 21 A. The Oocyst ........................................................................................................................ 21 B. Sporozoites ....................................................................................................................... 22 C. Attachment for Feeder Organelle ..................................................................................... 23 D. Trophozoites and Merozoites............................................................................................ 24 E. Sexual Stages .................................................................................................................... 25 VII. Transmission................................................................................................................................... 26 VIII. Prevention, Control, and Treatment ............................................................................................... 28 A. Physical Factors That Reduce Oocyst Viability ............................................................... 28 B. Chemicals That Reduce Oocyst Viability......................................................................... 31 C. Environmental Factors That Potentially Reduce Oocyst Numbers.................................. 33 References............................................................................................................................. 35
I.
Introduction
The genus Cryptosporidium is composed of protozoan parasites that infect epithelial cells in the microvillus border of the gastrointestinal tract of all classes of vertebrates. They are found worldwide. Effects of infection vary with the species of Cryptosporidium. Some species of Cryptosporidium infect many host species, whereas others appear restricted to groups such as rodents or ruminants, and still others are known to infect only one host species. Some species primarily infect the stomach, whereas others primarily infect the intestine. Some species are pathogenic, whereas the presence of others has not been shown to be related to any disease manifestations. Some infections are acute and self-limiting, whereas others are chronic. The severity and duration of infection with pathogenic species are also affected by 1
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2
Cryptosporidium and Cryptosporidiosis, Second Edition
the immune status of the infected person or animal. Immunocompetent individuals might suffer mild, moderate, or severe acute illness, whereas immunocompromised individuals can suffer severe chronic illness and even death. There are now 16 recognized species, and nearly triple this number of unnamed cryptosporidians have been identified only as genotypes. However, most studies on the biology, morphology, biochemistry, host preferences, immunology, pathogenicity, physiology, and prevalence have been conducted on one species, Cryptosporidium parvum. The reasons: this species is of medical and veterinary importance; it is geographically widespread; it infects many host species, producing prodigious numbers of oocysts, thus making it more easily obtainable for study than other species of Cryptosporidium; and it can be grown and tested in vitro and in animal models. Consequently, data derived from Cryptosporidium parvum, in some cases, have become generalized and extended to other members of the genus. Obviously, each species and genotype has individual characteristics that make it different from the others. It is important to keep this in mind when considering information based largely on one or a few species.
II.
History
Ernest Edward Tyzzer was the first person to recognize, clearly describe, and publish an account of a parasite he frequently found in the gastric glands of a tame variety of common mice. In 1907, he described its asexual and sexual stages (Tyzzer, 1907). Each had an organelle resembling the epimerite of gregarines that was specialized for attachment to host cells. He also noted that spores (oocysts) were excreted in the feces. He identified the parasite as a sporozoan of uncertain taxonomic status and named it Cryptosporidium muris. In 1910, he described the parasite in greater detail, again in the gastric glands of tame varieties of the common mouse, and proposed the name Cryptosporidium as a new genus and C. muris as the type species. He added Japanese waltzing mice and English mice as hosts and speculated that sporozoites from oocysts in the gastric glands might autoinfect these hosts (Tyzzer, 1910). With the exception that Tyzzer thought the endogenous developmental stages were extracellular because of the limitations of his compound microscope, all other aspects of his original description of the C. muris life cycle have been confirmed by electron microscopy. He noted that “it appears reasonably certain that the organisms observed in the gastric glands” of mice several years earlier by J. Jackson Clark (1894–1895) were the same species he was about to describe and not a parasite named Coccidium falciforme as Jackson supposed. In 1912, Tyzzer described another new species, Cryptosporidium parvum (Tyzzer, 1912). He demonstrated that C. parvum developed only in the small intestine of experimentally infected tame laboratory mice and that the oocysts of C. parvum were smaller than those of C. muris. He remained ambiguous on the subject of whether stages were intracellular or extracellular and noted that stages similar to those of C. parvum in the mouse were present in the small intestine of the rabbit. In 1929, Tyzzer illustrated, but did not clearly describe, the developmental stages of what he believed to be C. parvum in the cecal epithelium of chickens (Tyzzer, 1929). This parasite was later named C. tyzzeri in his honor, but the description lacked useful taxonomic data and is considered a nomen nudum. In the late 1920s and 1930s Tyzzer was considered the leading researcher on a related genus of economically important parasites, Eimeria. He described the new species E. acervulina, E. maxima, and E. mitis that caused coccidiosis in chickens and the new species E. meleagridis, E. dispersa, and E. meleagrimitis that infect turkeys. In 1934, Tyzzer was elected President of the American Society of Parasitologists. For 48 years after Tyzzer’s first publication, because Cryptosporidium was not recognized to be of economic, medical, or veterinary importance, there were no subsequent studies and therefore no new information on the genus. Two new species were named, but these were misidentified organisms that were not even in the genus Cryptosporidium (Wetzel, 1938; Matschoultsky, 1947). In 1955, after the report of a valid new species in turkeys, Cryptosporidium meleagridis, the first species of Cryptosporidium associated with illness and death (Slavin, 1955), interest still remained low. In the early 1970s, only slightly more interest was aroused when Cryptosporidium was reported to be associated with diarrhea in cattle (Panciera et al., 1971; Meutin et al., 1974) (see Figure 1.1).
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General Biology
3
FIGURE 1.1 A “Fault Tree” depicting sources of oocysts and routes of transmission. (Courtesy of Nancy Fayer.) (From Fayer, R., 1997. Cryptosporidium and Cryptosporidiosis. CRC Press, Boca Raton, FL. With permission.)
In 1976, two groups separately reported the first two cases of cryptosporidiosis in humans (Nime et al., 1976; Meisel et al., 1976). A 3-year-old child and a 39-year-old immunosuppressed patient shared the following backgrounds: each lived on a farm with cattle present, had a dog, presented with severe watery diarrhea, and was diagnosed by microscopic examination of an intestinal biopsy specimen. A third human case of enteritis was reported in a 9-year-old boy with congenital hypogammaglobulinemia who had no unusual exposure to animals or house pets (Lasser et al., 1979). The fourth case reported in the 1970s involved a 52-year-old, IgA-deficient, immunosuppressed renal transplant patient who had no known contact with animals (Weisburger et al., 1979). In addition to these first four human cases, reports of cryptosporidiosis during this decade appeared in nearly 40 publications on cattle, sheep, pigs, horses, turkeys, rabbits, monkeys, snakes, and guinea pigs. In the first four score years of the 1900s, less than 100 reports had been published on cryptosporidiosis, but by the last year of the 1980s, there were nearly 1200 reports. This surge of worldwide interest and study in the genus Cryptosporidium followed a report in 1982 from the Centers for Disease Control (CDC) describing 21 males from six large cities in the United States who had severe protracted diarrhea caused by Cryptosporidium in association with Acquired Immune Deficiency Syndrome (AIDS) (Anonymous, 1982). Approximately half of the new reports described the disease and pathology in humans and animals or addressed diagnosis and detection methods. In 1993, interest again increased dramatically following a massive waterborne outbreak in Milwaukee, Wisconsin, involving an estimated 403,000 persons (MacKenzie et al., 1994). The general public, public health agencies, agricultural groups, environmental groups, suppliers of drinking water, and others expressed concern and initiated studies on the basic biology of Cryptosporidium with emphasis on developing methods for recovery and detection of the oocyst stage, and prevention and treatment of the disease. In the mid-to-late 1990s, molecular methods for detection and identification emerged, were refined, and became the new cornerstone for taxonomic and epidemiologic studies. Subtyping, and
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Cryptosporidium and Cryptosporidiosis, Second Edition TABLE 1.1 Taxonomic Classification of Cryptosporidium Classification
Name
Empire
Eukaryota
Kingdom
Protozoa Goldfuss 1818 Apicomplexa Levine, 1970
Phylum
Class
Coccidea Leuckart, 1879
Order
Eucoccidiorida Leger and Duboscq, 1910 Cryptosporidiidae Leger, 1911
Family
Genus Type Species
Cryptosporidium Tyzzer, 1910 Cryptosporidium muris Tyzzer, 1910
Biological Characteristics Cells have nucleus containing most of cell’s DNA, enclosed by doublelayer membrane; nearly all have mitochondria. Predominantly unicellular; most have cristate mitochondria, Golgi bodies, and peroxisomes. Unicellular endosymbiont or predator with apical complex typically composed of polar rings, rhoptries, micronemes, and usually a conoid; subpellicular microtubules and micropore common. Oocyst generally contains infective sporozoites that result from sporogony. Reproduction both asexual and sexual. Locomotion by flexing, gliding, or undulation. Merogony; infects vertebrates and invertebrates. All development intracellular, extracytoplasmic, with feeder organelle; contains a single genus; found in all classes of vertebrates; life cycles are homoxenous; oocysts contain four naked sporozoites; after ingestion, sporozoites exit through a suture in oocyst wall; two or three types of meronts, the first generation possibly capable of recycling, the last forms microgametocytes and macrogametes; once fertilized, the zygote produces an oocyst; sporogony is endogenous. Sporocyst absent or united with oocyst so that the entire organism becomes a single spore with four sporozoites. Habitat, gastric glands of common mouse; schizont after nuclear division gives rise to 8 merozoites; early microgametocyte resembles schizont and gives rise to 16 microgametes 1.5 to 2.0 μm long; after fertilization, macrogamete develops thick cyst membrane, protoplasm divides and forms 4 sporozoites; mature oocysts ~ 7 × 5 μm; mature sporozoites slender and 12 to14 μm long; all development is extracellular; probable autoinfection by sporozoites in stomach.
Note: Portions of biological descriptions adapted from Tyzzer (1910), Levine (1985), and Upton (2000).
genomic, proteomic, and biochemical investigations followed. Of over 2300 publications in this decade, one-fourth addressed waterborne aspects of transmission or molecular findings. In the new millennium years approaching the centennial anniversary of the discovery and naming of Cryptosporidium, the emphasis on molecular and genetic studies as well as waterborne aspects of transmission continued.
III. Taxonomy All members of the genus Cryptosporidium are intracellular parasites. They are eukaryotic protozoa, which means that most of their DNA is contained within a nucleus surrounded by a double membrane, and they are unicellular organisms (Table 1.1). The genus Cryptosporidium is one of over 300 genera that include 4800 named species in the phylum Apicomplexa, but there may be ten times that number of species yet to be described. All are parasitic, and some are important disease agents such as malarial parasites of humans and animals, piroplasms, and coccidia (including Isospora, Toxoplasma, and Sarcocystis) of humans, coccidia (primarily Eimeria) of domesticated and wild animals, and gregarines of invertebrates. Over 150 species of mammals including humans (Table 1.2), as well as birds, reptiles, amphibians, and fish are parasitized by members of the genus Cryptosporidium. Most apicomplexans (except the eugregarines) have a complicated life cycle consisting of asexual and sexual stages resulting in sporogony. Specialized organelles found in the motile invasive asexual stages include subpellicular microtubules, important for locomotion, and an apical complex consisting of apical
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General Biology
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TABLE 1.2 Mammalian Host Species Infected with Cryptosporidium Order, Artiodactyla Addax nasomaculatus (Addax) Aepyceros melampus (Impala) Ammotragus lervia (Barbary sheep) Antidorcas marsupialis (Springbok) Antilope cervicapra (Blackbuck) Axis axis (Axis deer) Bison bison (American bison) Bison bonasus (European bison) Bos indicus (Zebu) Bos taurus (Ox) Boselaphus tragocamelus (Nilgai) Bubalus bubalis (Water buffalo) Bubalus depressicornis (Lowland anoa) Camelus bactrianus (Bactrian camel) Capra falconeri (Turkomen markhor) Capra hircus (Goat) Capreolus capreolus (Roe deer) Cervus albirostris (Thorold’s deer) Cervus duvauceli (Barasingha deer) Cervus elaphus (Red deer/elk/wapiti) Cervus eldi (Eld’s deer) Cervus nippon (Sika deer) Cervus unicolor (Sambar) Connochaetes gnou (Wildebeest) Connochaetes taurinus (Blue-eared gnu) Dama dama (Fallow deer) Elaphurus davidianus (Pere David’s deer) Gazella dama (Addra gazelle) Gazella dorcas (Dorca’s gazelle) Gazella leptoceros (Slender-horned gazelle) Gazella subgutterosa (Persian gazelle) Gazella thomsoni (Thomson’s gazelle) Giraffa camelopardalis (Giraffe) Hexaprotodom liberiensis (Pygmy hippopatomus) Hippotragus niger (Sable antelope) Kobus ellipsiprymmus (Ellipsen waterbuck) Lama glama (Llama) Lama guanicoe (Guanaco) Lama pacos (Alpaca) Muntiacus reevesi (Muntjac deer) Odocoileus hemionus (Mule deer) Odocoileus virginianus (White-tailed deer) Oryx gazella callotys (Fringe-eared oryx) Oryx gazella dammah (Scimitar horned oryx) Ovis aries (Sheep) Ovis musimon (Mouflon) Ovis orientalis (Urial) Sus scrofa (Pig) Syncerus caffer (African buffalo) Taurotragus oryx (Eland) Tayassu tajacu (Collared peccary) Tragelaphus eurycerus (Bongo) Order, Carnivora Acironyx jubatus (Cheetah) Canis familiaris (Dog) Canis latrans (Coyote)
Felis catus (Cat) Helarctos malayanus (Malayan bear) Martes foina (Beech marten) Meles meles (Badger) Mephitis mephitis (Striped skunk) Mustela putorius (Ferret) Panthera pardus (Leopard) Procyon lotor (Raccoon) Urocyon cinereoargenteus (Grey fox) Ursus americanus (Black bear) Ursus arctos (Brown bear) Ursus (Thalarctos) maritimus (Polar bear) Vulpes vulpes (Red fox) Zalophus californianus (California sea lion) Order, Chiroptera Eptesicus fuscus (Big brown bat) Myotis adversus (Large-footed mouse-eared bat) Order, Insectivora Ateletrix albiventris (African hedgehog) Crocidura russula (Greater white-toothed shrew) Erinaceus europaeus (European hedgehog) Sorex araneus (Long-tailed shrew) Sorex minutus (Pygmy shrew) Order, Lagomorpha Oryctolagus cuniculus (Rabbit) Sylvilagus floridanus (Cottontail) Order, Marsupialia Antechinus stuartii (Brown antechinus) Didelphis virginiana (Opossum) Isodon obesulus (Southern brown bandicoot) Macropus giganteus (Eastern grey kangaroo) Macropus rufogriseus (Red neck wallaby) Macropus rufus (Red kangaroo) Phascolarctos cinereus (Koala) Thylogale billardierii (Pademelon) Trichosurus vulpecula (Brushtail possum) Order, Perissodactyla Ceratotherium simum (Southern white rhinoceros) Equus caballus (Horse) Equus przewalski (Miniature horse) Equus zebra (Zebra) Rhinoceros unicornis (Rhinoceros) Tapirus terrestris (Brazilian tapir) Order, Primates Ateles belzebuth (Marimonda spider monkey) Calithrix jacchus (Common marmoset) Cercopithecus campbelli (Campbell’s mona) Cercopithecus talapoin (Talapoin monkey) Cercocebus albigena (Mangabey) Cercocebus torquatus (White-collared monkey) Cercopithecus aethiops (Velvet monkey) Erythrocebus patas (Patas monkey)
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Cryptosporidium and Cryptosporidiosis, Second Edition TABLE 1.2 (CONTINUED) Mammalian Host Species Infected with Cryptosporidium Eulemur macaco (Black lemur) Gorilla gorilla (Gorilla) Homo sapiens (Humans) Hylobates syndactylus syndactylus (Siamang) Lemur catta (Ring-tailed lemur) Lemur macacomayottensis (Brown lemur) Lemur variegatus (Ruffed lemur) Macaca fascicularis (Long-tailed macaque) Macaca fuscata (Japanese macaque) Macaca mulatta (Rhesus monkey) Macaca nemestrina (Cotton-tipped/pigtail macaque) Macaca radiata (Bonnet macaque) Macaca thibetana (Pere David’s macaque) Mandrillus leucophaeus (Drill) Nycticebus pygmaeus (Lesser slow loris) Papio anubis (Olive baboon) Papio cynocephalus (Baboon) Pithecia pithecia (White-faced saki) Pongo pygmaeus (Orangutan) Saguinus oedipus (Cotton-topped tamarin) Saimiri sciureus (Squirrel monkey) Varecia variegata (Red-ruffed lemur) Order, Rodentia Apodemus agrarius (Field mouse) Apodemus flavicollis (Field mouse) Apodemus sylvaticus (Field mouse) Castor canadensis (Beaver) Castor fiber (European beaver) Cavia porcellus (Guinea pig) Chinchilla laniger (Chinchilla) Clethrionomys glareolus (Red-backed vole)
Geomys bursarius (Pocket gopher) Glaucomys volans (Flying squirrel) Hystrix indica (Indian porcupine) Marmota monax (Woodchuck) Mesocricetus auratus (Golden hamster) Microtus agrestis (Field vole) Microtus arvalis (Orkney vole) Mus musculus (House mouse) Mus spretus (Western Mediterranean mouse) Myocastor coypus (Coypu) Ondatra zibethicus (Muskrat) Rattus norvegicus (Norwegian rat) Rattus rattus (House rat) Sciurus carolinensis (Gray squirrel) Sciurus niger (Fox squirrel) Sigmodon hispidus (Cotton rat) Spermophilus beecheyi (California ground squirrel) Spermophilus tridecemlineatus (13-lined ground squirrel) Tamias sibiricus (Siberian chipmunk) Tamias striatus (Chipmunk) Order, Monotremata Tacyglossus aculeatus (Echidna) Order, Proboscidea Elephas maximus (Indian elephant) Loxodonta africana (African elephant) Order, Sirenia Dugong dugon (Dugong) Phoca hispida (Ringed seal) Zalophus californianus (California sea lion)
rings, a conoid, micronemes, rhoptries, and dense granules, important for host cell invasion. Those genera that develop in the gastrointestinal tract of vertebrates and pass through all stages of their life cycle within a single host include Eimeria, Isospora, Cyclospora, and Cryptosporidium. These are often referred to as coccidia. Other coccidia that develop in the gastrointestinal tract of vertebrates and additionally are capable of or require extraintestinal development are referred to as tissue cyst-forming coccidia. These include Besnoitia, Caryospora, Frenkelia, Hammondia, Neospora, Sarcocystis, and Toxoplasma. Except for Toxoplasma, members of this latter group require two hosts—a predator and a prey—to complete the life cycle, and are termed obligatory heteroxenous parasites. Despite many morphological and life-cycle characteristics shared by Cryptosporidium species and all the aforementioned coccidia, some molecular analyses have suggested a closer relationship between Cryptosporidium species and gregarines (Carreno et al., 1999). Although phylogenetic analysis has placed cryptosporidians together with the aseptate archigregarines, it is not possible to determine if they have evolved from or are sisters to the gregarine clade (Leander et al., 2003).
A.
Requirements for Species Identification
The genus Cryptosporidium is characterized by specific morphologic and biological features (Table 1.1), but difficulty has arisen with characterization at the species level. This has resulted in part because, classically, many species of coccidia were identified and differentiated entirely on the basis of their unique oocyst morphology. This characteristic is not applicable within the genus Cryptosporidium because oocysts of many species lack unique features and are indistinguishable from one another. Because
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oocyst morphology alone is not sufficient to characterize a species of Cryptosporidium, other distinguishing factors must be included in the taxonomic description. Morphology of other life-cycle stages and variations within the life cycle (e.g., additional stages of merogony) can provide species-specific information. However, the need to examine every host for infected tissues that specifically demonstrate these characteristics renders them impractical. Determining the susceptibility of a host to infection by a species through infectivity studies can be helpful, but this trait also is encumbered by several potential problems. Variations in oocyst dosage, age, storage conditions, and processing methods, as well as the age, size, immune status, and genetic makeup of the host, can affect the outcome. Host specificity, however, can be extremely useful in supporting morphologic and genetic data when first establishing the species taxon, and knowledge of the range of potential hosts becomes important when studying the epidemiology of an outbreak or tracing possible sources of an infection. Biochemical-based methods have been used to identify differences within and between species. For example, isozyme analyses and two-dimensional gel electrophoresis patterns of surface proteins have provided detailed information. However, possibly, these techniques could produce identical zymograms or other gel electrophoresis banding patterns for multiple species or genotypes. Too few specimens have been tested to know which, if any, of these banding patterns are unique. Perhaps the overriding factor limiting the application of these methods is that each test requires very large numbers of noncontaminated oocysts, typically, 107 or more, a quantity often difficult or impossible to obtain. More recently, gene sequence information has become the most widely applicable factor for defining new species. In addition to species-specific unique data, the requirement for relatively small quantities of oocysts, and a relatively rapid processing time, there has been good correlation with the natural host range and host infectivity studies. Such genetic data have been based primarily on slight differences in the sequence of base-pairs within the gene, referred to as the 18s or small subunit ribosomal (ssr) RNA gene. The 18s sequence data have been included in the definition of new species, including C. andersoni, C. bovis, C. canis, C. galli, C. hominis, and C. suis. Additional genetic data have been supplied for the actin and HSP-70 genes, among others (Chapter 5). Confusion or uncertainty has arisen regarding how to distinguish interspecies differences from intraspecies variations. Often, this uncertainty has led to the naming of an isolate as a Cryptosporidium genotype (Table 1.3). Identifying an isolate or group of organisms within this genus as a genotype recognizes the incomplete knowledge of this isolate while also recognizing its uniqueness. Cryptosporidium genotypes are named when significant sequence differences in the small subunit rRNA or other genes are identified, and phylogenetic analysis has eliminated the possibility these differences are due to heterogeneity between copies of the gene or intragenotypic variations. A genotype is not a taxon. It is a partial and temporary descriptor. A taxon designation can be made with some confidence when substantial data become available. At the 6th Meeting on Molecular Epidemiology and Evolutionary Genetics in Infectious Disease, a symposium entitled “The taxonomy of the genus Cryptosporidium” was held, where the proposal was made that, when naming new species of Cryptosporidium, the following should be fulfilled: (1) provide morphometric data on oocysts; (2) provide genetic characterization; (3) demonstrate natural and, when feasible, experimental host specificity; and (4) comply with ICZN rules. Subgenotypes have also been recognized. These reflect differences within a genotype based on data obtained from GP 60 gene sequences (Chapter 5). Such differences help to distinguish sources of the parasite and its host range.
1.
Valid and Nonvalid Species
There are 16 valid named species of Cryptosporidium (Table 1.4), but undoubtedly, there will be many more. There are over 40 Cryptosporidium isolates referred to as genotypes (Table 1.3). These have been identified only as Cryptosporidium sp. followed by the genotype designation that reflects the host of origin for the oocyst stage. Some isolates, once identified as C. parvum or genotypes of C. parvum, have been elevated to species level, such as C. hominis, C. bovis, and C. suis. Other isolates of C. parvum, as well as those of C. canis, C. galli, and C. muris, have been differentiated at the molecular level and designated as genotypes or subgenotypes, and still others have been reported to vary in virulence.
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Cryptosporidium and Cryptosporidiosis, Second Edition TABLE 1.3 Genotypes of Cryptosporidium Intestinal C. C. C. C. C. C. C. C. C. C. C. C. C. C. C. C. C. C. C. C. C. C. C. C. C. C.
sp. Bear sp. Cervine 1, 2, 3 sp. Deer sp. Deer-like sp. Deer mice sp. Duck sp. Ferret sp. Fox and Fox II sp. Goose I and II sp. Horse sp. Marsupial I and II sp. Mongoose sp. Monkey sp. Mouse sp. Muskrat I and II sp. Opossum I and II sp. Ostrich sp. Ovine sp. Pig II (Pig I syn. C. suis) sp. Rabbit sp. Raccoon sp. Seal 1 and 2 sp. Sheep novel genotype (Chalmers et al.,) C. sp. Sheep novel genotype (Ryan et al.,) sp. Skunk sp. Snake canis Coyote; Dog; Fox
Gastric C. C. C. C. C. C.
sp. Caribou sp. Lizard sp. Tortoise sp. Woodcock galli Finch muris Japanese field mouse genotype
Additional traits of great enough significance to merit the relegation of these organisms to higher taxon status await discovery and documentation. Several species of Cryptosporidium were named, but subsequent morphologic data, cross-transmission studies, or reexamination of the descriptions has resulted in invalidating these names. Cryptosporidium ameivae, C. anserinum, C. nasorum, and C. tyzzeri lack enough description to properly identify them and are therefore considered nomen nuda. Cryptosporidium crotali, C. ctenosauris, C. lampropeltis, and C. vulpis were misidentified and are actually species of Sarcocystis. Cryptosporidium saurophilum, named for oocysts from a lizard after the name C. varanii, appeared in a somewhat obscure journal 3 years earlier. Cryptosporidium agni, C. cuniculus, C. garnhami, and C. rhesi are synonyms of C. parvum. Cryptosporidium curyi (cat source) with oocysts five to six times larger than those of C. parvum is a doubtful and unconfirmed species. Cryptosporidium bovis and C. suis originally lacked adequate descriptions and were considered synonyms of C. parvum until molecular data and morphological information were obtained and host transmission studies were conducted, establishing them first as genotypes of Cryptosporidium and later as valid species. Those species considered nonvalid, primarily because they were mistakenly identified and have been found to have oocysts or sporocysts characteristic of other genera, or the published descriptions lacked sufficient information for them to be clearly identified, or
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TABLE 1.4 Valid Named Species of Cryptosporidium, Type Host, and Oocysts Measurements Species
μm) Mean Oocyst Size/ Range (μ per Original Description
Author
Type Host
andersoni baileyi bovis canis felis galli hominis meleagridis molnari
Lindsay et al., 2000 Current et al., 1986 Fayer et al., 2005 Fayer et al., 2001 Iseki, 1979 Pavlasek, 1999 Morgan-Ryan et al., 2002 Slavin, 1955 Alvarez-Pellitero and Sitja-Bobadilla, 2002
7.4 6.2 4.9 5.0 5.0 8.3 5.2 5.2 4.7
C. muris C. parvum C. scophthalmi
Tyzzer, 1910 Tyzzer, 1912 Alvarez-Pellitero et al., 2004 Levine, 1980 (Brownstein et al., 1977)
Bos taurus (domestic cattle) Gallus gallus (chicken) Bos taurus (domestic cattle) Canis familiaris (domestic dog) Felis catis (domestic cat) Gallus gallus (chicken) Homo sapiens (human) Meleagris gallopavo (turkey) Sparus aurata (gilthead seabream) Dicentrarchus labrax (European seabass) Mus musculus (house mouse) Mus musculus (house mouse) Scophthalmi maximus (turbot) Elaphe guttata (corn snake)
2.8 to 3.6
C. C. C. C. C. C. C. C. C.
C. serpentis
C. suis C. varanii
Ryan et al., 2004 Pavalasek et al., 1995
C. wrairi
Vetterling et al.,1971
E. subocularis (rat snake) Sanzinia madagascarensus (Madagascar boa) Sus scrofa (domestic pig) Varanus prasinus (Emerald monitor) Cavia porcellus (guinea pig)
× × × × × × × × ×
5.5/6.0–8.1 4.6/5.6–6.3 4.6/4.8–5.4 4.7/3.7–5.9 4.5 6.3/8.0–8.5 4.9/4.4–5.9 4.6/4.5–6.0 4.5/3.2–5.5
× × × ×
5.0–6.5 4.5–4.8 4.2–4.8 3.7–5.9
× × × ×
6.2–6.4 4.4–5.4 4.2–5.3 3.0–5.0
7×5 ovoid or spherical ≥4.5 4.4 × 3.9/3.7–5.0 × 3.0–4.7
4.6 × 4.2/4.4–4.9 × 4.0–4.3 4.8 × 4.7/4.8–5.1 × 4.4–4.8 5.4 × 4.6/4.8–5.6 × 4.0–5.0
they were named without conforming to the rules of the International Code of Zoological Nomenclature (ICZN), are found in Table 1.5. Reexamination of the species name C. parvum has led to questions regarding its true identity. In 2006, in an attempt to clarify species classification, Slapeta proposed that the name C. pestis replace C. parvum for the zoonotic species prevalent in cattle that became the default species for many subsequent studies on human and mammalian cryptosporidiosis. The parasite Tyzzer found in laboratory mice and named C. parvum could be the organism now identified as the Cryptosporidium mouse genotype because Tyzzer could easily infect adult mice with his isolate, whereas the C. parvum widely recognized today produces very low-level infections in adult mice. The species or genotype Tyzzer described can never be positively identified because no recoverable specimens remain from Tyzzer’s studies. The name C. parvum was essentially validated for the zoonotic bovine isolate after a modern morphologic description of the oocysts as well as the life cycle and cross-transmission studies between mice and cattle were published (Upton and Current, 1985; Current and Reese, 1986). Rejecting these modern studies and establishing a new name for the zoonotic C. parvum bovine isolate would be prohibited by Article 23.9.1.2 of the ICZN, which states that prevailing usage must be maintained when “the junior synonym or homonym has been used for a particular taxon, as its presumed valid name, in at least 25 works, published by at least 10 authors in the immediately preceding 50 years and encompassing a span of not less than 10 years.” Although the species described by Tyzzer in 1912 was most likely not the C. parvum (bovine genotype) widely recognized today but more likely the mouse genotype that is biologically and genetically distinct, the mouse genotype will quite probably be thoroughly characterized and given a new species name.
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Cryptosporidium and Cryptosporidiosis, Second Edition
TABLE 1.5 Nonvalid Species Names Name C. agni
Author
Host Ovis aries (sheep)
C. anserinum
Barker and Carbonell, 1974 Arcay de Peraza and Bastardo de San Jose, 1969 Proctor and Kemp, 1974
C. baikalika
Matschoulsky, 1947
Scolopax sp. (woodcock)
C. crotalis
Triffit, 1925
Crotalus sp. (rattlesnake)
C. ctenosauris
Duszynski, 1969
Ctenosaura similis (lizard)
C. cuniculus
Inman and Takeuchi, 1979
Oryctolagus cuniculus (rabbit)
C. enteriditis
Qadripur and Klose, 1985
Homo sapiens (human)
C. garnhami
Bird, 1981
Homo sapiens (human)
C. lampropeltis
Anderson et al., 1968
C. nasorum
Hoover et al., 1981
Lampropeltis calligaster (kingsnake) Naso lituratus (tropical fish)
C. pestis C. rhesi
Slapeta, 2006 Levine, 1980
Bos taurus (cattle) Macaca mulatta (rhesus monkey)
C. saurophilum C. tyzzeri
Koudela and Modry, 1998 Levine, 1961
Eumeces schneideri (skunk) Gallus gallus (chicken)
C. villithecum
Paperna et al., 1986
Cichlid fish
C. vulpis
Wetzel, 1938
Vulpes vulpis (fox)
C. ameivae
a
Ameiva ameiva (lizard)
Anser anser (goose)
Reason Name Is Not Valid No oocyst measurements, no taxonomically useful data No oocyst measurements, no taxonomically useful data No oocyst measurements, no taxonomically useful data Gregarine oocyst misidentified as Crypto oocyst Sarcocystis sporocyst misidentified as Crypto oocyst Sarcocystis sporocyst misidentified as Crypto oocyst No oocyst measurements, no taxonomically useful data No oocyst measurements, no taxonomically useful data No oocyst measurements, no taxonomically useful data Sarcocystis sporocyst misidentified as Crypto oocyst No oocyst measurements, no taxonomically useful data Did not follow ICZNa rules No oocyst measurements, no taxonomically useful data Synonym of C. varanii No oocyst measurements, no taxonomically useful data Not a peer-reviewed article, insufficient data Sarcocystis sporocyst misidentified as Crypto oocyst
International Code of Zoological Nomenclature.
IV.
Host Specificity
To determine the host range for a species or genotype of Cryptosporidium, oocysts are obtained from animals of one species and fed to or intubated into animals of another species. If the life cycle is completed in the putative additional host species and oocysts are excreted that are genetically identical to those that initiated the infection, the host range is extended. Generally, isolates from one class of vertebrates have not been infectious for animals of another class. However, the host diversity already known among some species makes it impossible to establish this as a genus-wide paradigm (Table 1.6A, B). The avian parasites C. baileyi and C. meleagridis have been identified in and in both immunocompromised and healthy humans (Ditrich et al., 1991; Pedraza-Diaz et al., 2000; Guyot et al., 2001; Xiao et al., 2001; Gatei et al., 2003). Several subtypes of C. meleagridis have been described (Glaberman et al., 2001). The mammalian parasite C. parvum does not infect ducks or geese but does infect chickens. Conflicting reports add to the confusion regarding the range of susceptible host species for C. parvum. Whereas one study indicated that C. parvum oocysts from a human-infected fish, amphibians, reptiles, birds, and mammals based on finding oocysts in the feces of the recipients beginning 4 to 9 days after oral inoculation (Arcay et al., 1995), another found that C. parvum oocysts from a bovine did not infect
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TABLE 1.6A Major and Minor Species of Cryptosporidium Infecting Humans and Selected Domesticated Animals and Wildlife Host Camel Cat Cattle
Major Species
Chicken Coyote Deer (Red) Deer (White tail) Dog Duck and goose Fish Fox Goat Guinea pig Horse Human
C. andersoni, C. parvum? C. felis C. parvum, C. bovis, C. andersoni deerlike genotype C. baileyi C. canis coyote genotype C. parvum C. parvum, deer genotype C. canis Goose genotype I and II C. scophthalmi, C. molnari C. canis fox genotype C. parvum C. wrairi Horse genotype C. hominis, C. parvum
Lizard Mouse Muskrat Pig Sheep Snake Squirrel Turkey
C. serpentis, C. varanii C. muris, mouse genotype muskrat genotype I C. suis Cervine genotypes 1–3, bovine genotype C. serpentis C. muris, squirrel genotype C. meleagridis, C. baileyi
Minor Species
C. suis C. meleagridis, C. galli
C. baileyi, duck genotype C. canis dog genotype, fox genotype II
C. meleagridis, C. felis, C. canis, C. suis, C. baileyi, cervine genotype Lizard genotype Muskrat genotype II Pig genotype II C. parvum, sheep novel genotypes C. varanii, snake genotype
fish, amphibians, or reptiles but simply passed through the digestive tract without infecting enterocytes (Graczyk et al., 1996). Even within a vertebrate class, oocysts of one species of Cryptosporidium do not always infect more than one host species. Cryptosporidium wrairi is known to infect only guinea pigs (Gibson and Wagner, 1986), and C. bovis to infect only cattle (Fayer et al., 2005). Other species of Cryptosporidium have been found to infect a predominant host species and, apparently with rare exceptions, additional or minor hosts. For example, C. felis infects cats (Iseki, 1979; Asahi et al., 1991), C. canis infects dogs (Fayer et al., 2001), C. andersoni infects cattle, C. muris infects mice, and C. suis (Ryan et al., 2004) infects pigs. However, C. felis, C. andersoni, C. canis, C. suis, and the Cryptosporidium cervine genotype were detected in 0.2, 0.1, 0.04, 0.04, and 0.04%, respectively, of 2414 humans with diarrhea (Leoni et al., 2006). In contrast, C. parvum has been thought to infect many, if not all, species of mammals.
V.
Life Cycle
A diagrammatic life cycle for Cryptosporidium spp. is shown in Figure 1.2. The primary site of infection with C. hominis and C. parvum is the small intestine. In some animals such as mice (Figure 1.17) and calves, the ileum above the cecal junction is the favored location for C. parvum. Cryptosporidium has been found in extraintestinal sites in animals (Fleta et al., 1995) and in some severely immunocompromised humans (Chapter 4). Other species such as C. muris, C. andersoni, and C. serpentis favor the gastric mucosa (Chapters 14, 16, and 18). Cryptosporidium baileyi, in chickens, favors the respiratory tree and the cloaca (Figure 1.18) (Chapter 15).
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Cryptosporidium and Cryptosporidiosis, Second Edition TABLE 1.6B Geographic Distribution of Reported Human Cryptosporidiosis Africa Algeria Burundi Cameroon Ethiopia Gabon Ghana Guinea Guinea-Bissau Ivory Coast Kenya Kwa-Zulu Natal Liberia Malawi Mali Mauritania Morocco Nigeria Rwanda Senegal South Africa Sudan Tanzania Togo Tunisia Uganda Zaire Zambia Zimbabwe
Europe Austria Belarus Belgium Czech Republic Denmark England Finland France Germany Greece Hungary Ireland Italy Lithuania Netherlands Poland Portugal Romania Russia Scotland Serbia Slovenia Spain Sweden Switzerland Turkey Wales North America
Pacific Australia Indonesia Malaysia New Zealand Papua-New Guinea Philippines Singapore
A.
Canada United States
Central/South America Argentina Bolivia Brazil Chile Colombia Costa Rica Ecuador El Salvador Guatemala Guyana Mexico Panama Peru Uruguay Venezuela Asia Bangladesh Cambodia China India Japan Korea Myanmar Nepal Pakistan Sri Lanka Taiwan Thailand Vietnam Caribbean
Middle East Azerbaijan Egypt Iran Israel Kuwait Saudi Arabia
Cuba Haiti Jamaica Puerto Rico St. Lucia Tobago Trinidad Virgin Islands
The Environmental Stage and Ultimate Source of Infection
The sporulated oocyst, the only documented exogenous stage, is excreted from the body of an infected host in the feces. It consists of a tough trilaminar wall that surrounds and maintains the viability of four internal sporozoites under adverse environmental conditions. Those sporozoites ultimately are the source of new infections.
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FIGURE 1.2 Diagrammatic representation of the life cycle of C. parvum. (Modified from Fayer, R., 1997. Cryptosporidium and Cryptosporidiosis. CRC Press, Boca Raton, FL. With permission.)
B.
Excystation
The endogenous phase begins after the oocyst is ingested by a suitable host. Once inside the body, the first step towards infection is excystation, the opening of the oocyst wall along a suture at one pole of the oocyst through which the four infectious sporozoites leave the oocyst (Figure 1.5). For most apicomplexan parasites related to Cryptosporidium, excystation of sporozoites requires that oocysts be exposed to reducing conditions in the stomach followed by exposure to pancreatic enzymes and/or bile salts in the small intestine. For Cryptosporidium, such exposure may enhance excystation, but under experimental conditions, it has been shown that sporozoites can excyst from oocysts in warm aqueous solutions alone, possibly enabling autoinfection and infections reported in extraintestinal sites such as the conjunctiva of the eye, the respiratory tract (Fayer et al., 1990), gall bladder, lymph nodes, testicle, ovary, uterus, and vagina (Fleta et al., 1995).
C.
Cell Invasion and Internalization
Sporozoites that excyst from oocysts are motile, approach a potential host-cell anterior (apical) end first, and actively invade the cell (Wetzel et al., 2005). They are characterized by the presence of secretory organelles that exocytose during the invasion process (Figure 1.3) (O’Hara et al., 2005). When the sporozoite initially contacts the host cell membrane, the single rhoptry extends to the attachment site, and the micronemes and dense granules move to the apical complex region (Huang et al., 2004). Sporozoites (and merozoites) of C. parvum express a 1300-kDa conserved apical complex glycoprotein called CSL that contains a ligand involved in attachment to intestinal epithelial cells during the infection process (Riggs et al., 1997; Langer and Riggs, 1999). Binding of CSL was found to be localized to a surface-exposed, conserved receptor on the microvillar surface of cells of epithelial origin, (Langer et
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FIGURE 1.3 Transmission electron micrographs (TEMs). Initial stages of cell invasion. (A) and (D) are serial images of a C. parvum sporozoite adjacent to a H69 human bile duct epithelial cell showing membrane protrusion. (B), (C), and (E) are high magnifications of the boxed areas of (A) and (D). Arrows and arrowheads in (C) and (D) show a dense band underlying the host cell membrane where the parasitophorous vacuole is forming at the parasite–host cell interface. Vacuolelike structures appear in the apical end of the sporozoite (E, asterisk). Vacuole-like structures in the apical complex of a sporozoite with a vacuole bubbled out toward the host cell suggest secretion from the parasite into the cell (F); these are shown at a higher magnification in (G). Bars in (A) and (D) = 0.5 μm, bars in (B), (C), (E), and (G) = 0.2 μm. (From Huang et al., 2004. J. Parasitol. 90, 212–221. With permission.)
al., 2001). Similar findings have been reported for additional mucin-like glycoproteins localized at the apical region (Cevallos et al., 2000) and for a thrombospondin-related adhesive protein crucial for invasion of host cells (Spano et al., 1998). The attached sporozoites, initially slender and crescent or boomerang shaped, become oval or spherical. Vacuoles form near the anterior end, possibly arising from the parasite, and cluster together, surrounding the parasite and forming a preparasitophorous vacuole (Figure 1.4) (Huang et al., 2004). This vacuolar area fuses with the host-cell membrane to form a host–parasite interface. Thin membrane-bound cytoplasmic extensions from the host cell microvilli surround the parasite, eventually containing it within a mature parasitophorous vacuole (Umemiya et
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FIGURE 1.4 TEMs of internalization of the sporozoite stage. (A), (C), and (E) are serial images of a C. parvum sporozoite becoming spherical and internalized in a H69 human bile duct epithelial cell. The preparasitophorous vacuole membrane extending from the host cell appears to move posteriad to the sporozoite as the host-cell membrane protrudes and fuses with it. (B), (D), and (F) are higher magnifications of the boxed areas of (A), (C), and (E), respectively. A tunnel (arrows) at the original site of attachment connects the vacuole-like apical complex region (asterisks) of the sporozoite with the hostcell cytoplasm. (From Huang et al., 2004. J. Parasitol. 90, 212–221. With permission.)
al., 2005). Sporozoites within the parasitophorous vacuole are intracellular but are not directly in contact with the host-cell cytoplasm—they are extracytoplasmic. During formation of the parasitophorous vacuole, material appears to be released from the conoid at the anterior tip of the sporozoite, while a large membrane-bound electron-lucent vacuole forms within the anterior third of the sporozoite where the rhoptry and micronemes had been present (Figure 1.4). Also during this time, vacuole-like structures at the apical end of the sporozoite appear to be directly connected to the host-cell cytoplasm. Both an electron-dense band beneath this site and the parasitophorous vacuole keep the parasite intracellular but extracytoplasmic. The electron-dense band subsequently matures into a unique structure referred to as
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FIGURE 1.5 Scanning electron micrograph (SEM) of oocysts and excysting sporozoites. (A) Intact oocyst before excystation. (Magnification ×16,000). (B) Three sporozoites (Sp) excysting from an oocyst simultaneously via the cleaved suture (Su) (Magnification ×16,000). (C) Empty oocyst (Magnification ×16,000). (D) Excysted sporozoite; Ae, apical end. (Magnification ×14,000). All figures originally from Reduker et al., 1985. J. Protozool. 32, 708. With permission; Also from Fayer, R., 1997. Cryptosporidium and Cryptosporidiosis. CRC Press, Boca Raton, FL. With permission.)
an attachment or feeder organelle. As this internalization process progresses, the sporozoite becomes spherical and is called a trophozoite (Figures 1.4, 1.5, 1.6, and 1.7). In cell culture, attachment and internalization was completed in 15 min (Lumb et al., 1988).
D.
Asexual Multiplication (Merogony)
Asexual multiplication called merogony (syn. schizogony) results when the trophozoite nucleus divides (Figure 1.8). Cryptosporidium baileyi has three types of meronts (syn. schizonts), and C. parvum has two types. For C. parvum, Type I meronts develop six or eight nuclei, each becomes incorporated into a merozoite, a stage structurally similar to the sporozoite (Figure 1.9). Each mature merozoite, theoretically, leaves the meront to infect another host cell (Figure 1.10) and to develop into another Type I or into a Type II meront. Type II meronts produce four merozoites.
E.
Sexual Reproduction (Gamogony)
It is thought that only merozoites from Type II meronts initiate sexual reproduction upon infecting new host cells by differentiating into either a microgamont (male) or a macrogamont (female) stage. Each microgamont (syn. microgametocyte) becomes multinucleate, and each nucleus is incorporated into a microgamete, a sperm-cell equivalent (Figure 1.11). Macrogamonts remain uninucleate, an ovum
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FIGURE 1.6 TEMs. (A) High magnification of apical complex of merozoite showing apical rings (Arl,2), micronemes (Mn), plasmalemma (Pl), inner membrane complex (Im), subpellicular microtubule (Sm), and electron-dense collar (Ec) (Magnification ×99,000). (B) Merozoite in early stage of attachment to an epithelial cell; Ai, anterior invagination; El, electron-dense layer of attachment zone; Mn, microneme; No, nucleolus; Nu, nucleus; Ph, plasmalemma of host cell; Pv, parasitophorous vacuole (Magnification ×32,500). (C) Slightly more advanced stage of attachment; note the enlarged anterior invagination (Ai), electron-dense collar (Ec), electron-dense layer (El), and fibrous layer (Fl) of attachment organelle (Magnification ×34,000). (D) Trophozoite with electron-dense collar located at margins of attachment organelle (Magnification ×19,500). (From Fayer, R., 1997. Cryptosporidium and Cryptosporidiosis. CRC Press, Boca Raton, FL. With permission.)
equivalent. How microgametes detect macrogamonts is unknown, but they must attach to and penetrate the host-cell membrane and the macrogamont cell membrane for fertilization to occur (Figure 1.12). After the microgamete enters the macrogamont cytoplasm, it either enters the nucleus or passes its nuclear contents through the nuclear membrane of the macrogamont (Figure 1.13). It is assumed that only fertilized macrogamonts develop into oocysts. In the transition, a trilaminar wall forms around the macrogamont, the 2N nucleus of the fertilized macrogamont undergoes meiosis, and four 1N sporozoites develop.
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FIGURE 1.7 (A) TEM of C. parvum sporozoite relict mitochondrion (asterisk) surrounded by a double membrane (arrows) posterior to the nucleus (N) and adjacent to the crystalloid body (CB) of unknown origin and function. (From Keithly et al., 2005. J. Eukaryot. Microbiol. 52, 132–140. With permission). (B) SEM of micronemes isolated from C. parvum sporozoites after sucrose density gradient. The rod-like, stain-excluding, smooth-surfaced, micronemes appear intact, and those with a quasi-helical internal organization appear damaged. (From Harris et al., 2003. Micron 43, 65–78. With permission.)
FIGURE 1.8 TEMs of schizogony. (A) Schizont showing early stages of merozoite formation at margin of schizont; note that a merozoite bud (Mb) is developing above each pole of the two nuclei (Nu) visible in this section; Rb, residual body (Magnification ×15,000). (B) Merozoites in an advanced stage of budding from the schizont residual body (Rb); Pv, parasitophorous vacuole (Magnification ×14,000). (From Fayer, R. 1997. Cryptosporidium and Cryptosporidiosis. CRC Press, Boca Raton, FL. With permission.)
F.
Sporogony
Oocysts sporulate in situ and, when mature, contain four sporozoites (Figure 1.14). Oocysts in the gastrointestinal tract are excreted with feces, whereas those in the respiratory tract exit the body with respiratory or nasal secretions. Some reports suggest that oocysts with thin walls release sporozoites that autoinfect the host, whereas those with thicker walls leave the body to infect other hosts (Current, 1985, 1988).
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FIGURE 1.9 TEM of mature schizont showing merozoites (Mz). Note dense granules (Dg), electron-dense collar (Ec), micronemes (Mn), nucleolus (No), nucleus (Nu), parasitophorous vacuole (Pv), and rhoptry (Rh) (Magnification ×28,000). (From Fayer, R. 1997. Cryptosporidium and Cryptosporidiosis. CRC Press, Boca Raton, FL. With permission.)
Cryptosporidium and Caryospora are the only coccidia in which oocysts sporulate in situ followed by autoinfection. Autoinfection results when sequential asexual and sexual phases of the life cycle are repeated within the same host, beginning with sporozoites released from the oocysts that have developed in situ after sporogony. Oocysts of the related coccidia, Eimeria, Hammondia, Isospora, and Toxoplasma, are excreted unsporulated and do not become infectious until they sporulate, forming sporozoites, outside the body. As in the case of Cryptosporidium, sporogony for Frenkelia and Sarcocystis takes place internally. However, their life cycles differ from that of Cryptosporidium. These genera are obligatorily heteroxenous, i.e., sexual stages and oocysts develop in carnivore (predator) hosts, and oocysts are infective only for the intermediate (prey) host species in which asexual multiplication takes place and tissue cysts develop.
G.
Prepatent Patent Times
The prepatent period is the shortest time after infective oocysts have been ingested, the endogenous life cycle has been completed, and the first newly developed oocysts have been excreted. This time varies with the host and species of Cryptosporidium as well as the infective dose. Experimentally determined prepatent periods range from 2 to 7 days for C. parvum in calves (Tzipori et al., 1983) and 4 to 22 days for humans (DuPont et al., 1995), 2 to 9 days for C. suis in pigs (Enemark et al., 2003), 10 to 12 days for C. bovis in cattle (Fayer et al., 2005), 6 to 21 days for C. muris in mice (Matsui et al., 1999), 5 to 6 days for C. felis in cats (Iseki, 1979), and 4 to 24 days for C. baileyi in chickens (Current et al., 1986). The patent period is the duration of infection usually characterized by the days of oocyst excretion. Experimentally determined patent periods range from 1 to 12 days for C. parvum in calves and 1 to 20
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FIGURE 1.10 SEM of cryptosporidia on sheep intestinal epithelial cells. Eight merozoites are escaping from one host cell, and the impressions of merozoites are visible in the host cell at the upper right (Magnification ×15,750). (From Fayer, R. 1997. Cryptosporidium and Cryptosporidiosis. CRC Press, Boca Raton, FL. With permission.)
days for humans, 9 to 15 days for C. suis in pigs, 18 days for C. bovis in cattle, 7 to 10 days for C. felis in cats, and up to 18 days for C. baileyi in chickens.
H.
Extracellular Stages
Gamont-like stages have been observed extracellularly in cell cultures infected with C. andersoni (Hijjawi et al., 2002) and in cell-free medium inoculated with C. parvum (Hijjawi et al., 2004). The morphological characteristics of these extracellular stages resembled those of gregarines, but molecular testing confirmed them to be C. andersoni (Hijjawi et al., 2002). Comparative observations were made of gamont-like extracellular stages from intestinal contents of mice infected with C. parvum and in in vitro cultures infected with C. parvum (Hijjawi et al., 2002). However, the origin and fate of these stages remain
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FIGURE 1.11 TEMs of microgamonts. (A) Microgamont with microgametes budding (Bm) from a residual body (Rb) (Magnification ×16,000). (B) Mature microgamont with portions of 6 microgametes (Mi) separated from the residual body (Rb) (Magnification ×19,000). (From Fayer, R. 1997. Cryptosporidium and Cryptosporidiosis. CRC Press, Boca Raton, FL. With permission.)
unclear, and their identity is controversial (Rosales et al., 2005; Girouard et al., 2006) pending further confirmation from independent sources.
VI. Structure A.
The Oocyst
The oocyst wall of Cryptosporidium parvum is a trilaminar structure with an average thickness of ~ 49 nm (Harris and Petry, 1999) (Figures 1.14 to 1.16). The outer layer is irregular in thickness but averages 10 nm (Reduker et al., 1985). Beneath this layer is a 2.5 nm-thick electron-lucent layer. Beneath this is the inner layer composed of two zones. The outer zone is 11.6 nm and the inner zone 25.8 nm thick. The wall is continuous except at one pole where it is interrupted by a single seam or suture that extends one-third to one-half the way around the periphery. When the suture opens during excystation, the oocyst wall on either side of the suture retracts, coiling inward, and sporozoites exit the oocyst (Figures 1.5 and 1.16). A trypsin-bile-sensitive suture similar in appearance to that of C. parvum is found in the sporocyst wall of Sarcocystis, Toxoplasma, Isospora, and Goussia, suggesting a close phylogenetic relationship between Cryptosporidium and the sarcocystids or the calyptosporids (i.e., Goussia, Calyptospora). The outer wall is thought to be a glycoprotein. The central layer of the wall is thought to be a glycolipid/lipoprotein that imparts some rigidity and is responsible for the acid-fast staining property of the wall. The inner layer, which appears linear and filamentous, is thought to be a glycoprotein, providing some rigidity and elasticity to the wall. Thin-walled oocysts differ in that the wall does not contain a thick multizoned inner layer. Contained within the oocyst is a membrane-enclosed residual body and four sporozoites. The residual body contains a large lipid body, many amylopectin granules, a crystalline protein inclusion, ribosomes, and cytomembranes (Figures 1.14 and 1.16). The amylopectin granules consist of complex branched polysaccharide chains of two morphologic types—larger smooth-surfaced granules with a coiled interior
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FIGURE 1.12 TEMs of microgametes. (A) Longitudinal section. Note concentric lamellae (Cl) just posterior to the apical cap (Ac); Eg, electron-dense granules; Mt, microtubule; Ni nucleus of microgamete; Pl, plasmalemma of microgamete (Magnification ×75,600). (B) Cross section showing microtubules (Mt) in close proximity to the nuclear envelope (Ne), microgamete nucleus (Ni), plasmalemma (Pl), and inner membrane (Im) of microgamete (Magnification ×66,000). (C) High magnification of a microgamete in close proximity to a host cell containing a macrogamont; plasmalemma of microgamete (single arrow) and macrogamont (double arrow); Ac, apical cap; Cl, concentric lamellae; M, nucleus of microgamete; Ph, plasmalemma of host cell; Pm, parasitophorous vacuolar membrane (Magnification ×80,000). (From Fayer, R. 1997. Cryptosporidium and Cryptosporidiosis. CRC Press, Boca Raton, FL. With permission.)
resembling a ball of string, and smaller granules with an irregular surface and a rodlike particulate interior (Harris et al., 2004).
B.
Sporozoites
Sporozoites of Cryptosporidium parvum appear similar to those of other coccidia. Transmission electron microscopy (TEM) has revealed the presence of organelles typical of the phylum (Figures 1.6 and 1.7). Beneath the surrounding pellicle are subpellicular microtubules. They originate at the electron-dense apical collar with associated apical rings and extend posteriad. The apical complex consists of the apical rings (the anteriormost structure), the conoid, and organelles for invading host cells: the single rhoptry, the micronemes, and electron-dense granules. Rhoptries of apicomplexans are flask-shaped, membranebound, electron-dense secretory organelles thought to originate from the endoplasmic reticulum by Golgi secretion (Sam-Yellowe, 1996). Micronemes in freshly excysted sporozoites appear stacked as rows of rod-like bodies. Each microneme, ~100 × 35 nm, has an internal quasi-helical, beaded pattern thought to be a protein polymer (Harris et al., 2003). The Golgi organelle, the nucleus, small amylopectin granules, and ribosomes are more centrally located. Posterior to the nucleus is an enigmatic crystalloid body for which neither the origin nor function is known (Figure 1.7). Between the crystalloid body and
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FIGURE 1.13 TEM of a macrogamont containing a microgamete nucleus (Ni); Am, amylopectin granules; Ec, electron-dense collar; Fo, feeding organelle; Go, Golgi complex; Na, nucleus of macrogamont; No, nucleolus of macrogamont; Pv, parasitophorous vacuole; Wf, wall-forming body (Magnification ×34,000). (From Fayer, R. 1997. Cryptosporidium and Cryptosporidiosis. CRC Press, Boca Raton, FL. With permission.)
the nucleus, and surrounded by multiple segments of rough endoplasmic reticulum, is a double-membrane-bound, mitochondrion-related organelle referred to as a relict mitochondrion (Figure 1.7) (Keithly et al., 2005). This unusual mitochondrion contains internal compartments but lacks the typical tubular cristae seen in other coccidia. It resembles the highly reduced nucleus of some endosymbiotic cryptomonad algal cells found within unicellular hosts (Putignani, 2005).
C.
Attachment for Feeder Organelle
Sporozoites and merozoites bound to epithelial cells by specific receptors become enveloped by the hostcell membrane until they become intracellular within a parasitophorous vacuole (Figures 1.3 and 1.4). Changes in the apex of the host cell and in the parasite result in the formation of an attachment or feeder organelle (Figures 1.13 and 1.14). At the contact site, an electron-dense layer forms in the host cell immediately above the terminal web of microfilaments, whereas the parasite plasmalemma invaginates at the apex just below the apical rings, which disappear (Figures 1.3 and 1.4). This invagination enlarges as an electron-dense collar expands laterally, eventually locating close to the lateral margin of the electron-dense layer. Concurrently, a fibrous layer that spans the area within the electron-dense collar forms between the invaginated parasite plasmalemma and the electron-dense layer of the host cell. The plasmalemma in this region becomes highly convoluted, resulting in a large surface area confined to a relatively small space. This site has been referred to as both the attachment organelle and the feeder organelle, the latter being the point of nutrient recruitment from the host cell. The organelle appears separable from the schizont residual body only after merozoites are fully formed. Microgametocytes
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FIGURE 1.14 TEMs of intracellular oocysts. (A) Thin-walled oocyst; Am, amylopectin; Fo, feeding (attachment) organelle; Lb, lipid body; Ow, thin oocyst wall; Pv, parasitophorous vacuole; Sp, sporozoite (Magnification ×23,240). (B) Thick-walled oocyst with suture (Su) at one pole; Lb, lipid body; Ow, thick oocyst wall; Pv, parasitophorous vacuole; Sp, sporozoite (Magnification ×23,240). (From Fayer, R. 1997. Cryptosporidium and Cryptosporidiosis. CRC Press, Boca Raton, FL. With permission.)
separate only after microgametes mature (Figure 1.11), and the mature zygote separates from the organelle after the oocyst sporulates in situ (Figure 1.14).
D.
Trophozoites and Merozoites
Trophozoites contain a prominent nucleolus within a single nucleus surrounded by cytoplasm (Figure 1.6), and a well-developed attachment/feeder organelle. During nuclear division of merogony, division spindles, nuclear plaques, and centrioles have not been observed. After nuclear division, merozoites develop simultaneously around the margin of the meront (syn. schizont). Merozoite anlagen, including a double inner membrane complex, an electron-dense collar, electron-dense granules, a few micronemes, ribosomes, and cytoplasm, form immediately beneath the schizont plasmalemma just above each schizont nucleus. Merozoites remain attached to a residuum at their posterior end and become more elongated
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FIGURE 1.15 High-magnification TEMs. (A) Margin of zygote showing two membranes (arrows) external to the parasite plasmalemma (Pl) and two types of wall forming bodies (Wfl and Wf2); Im, inner membrane of zygote (Magnification ×80,000). (B) Showing suture in oocyst wall (Magnification ×118,000). (C) Cross section of completely formed oocyst wall showing inner (Io) and outer (Oo) layers (Magnification ×118,000). (From Fayer, R. 1997. Cryptosporidium and Cryptosporidiosis. CRC Press, Boca Raton, FL. With permission.)
during maturation. During this time, a pair of rhoptries, more micronemes, numerous electron-dense granules, and many ribosomes form in the cytoplasm (Figure 1.9). Micronemes often form in rows perpendicular to the long axis of the merozoite (Figure 1.9). Upon maturity, merozoites separate from the residual body, the host-cell membrane surrounding the meront lyses, and merozoites become extracellular (Figure 1.10), able to infect other host cells. Numerous developmental stages including meronts containing fully formed merozoites can be seen in Figures 1.17 and 1.18.
E.
Sexual Stages
Microgamonts have been found less frequently than other stages. Immature microgamonts resemble meronts but contain small, compact nuclei. The single-surface membrane later doubles at sites around the margin where microgametes form (Figure 1.11). Each microgamete forms as a nuclear protrusion at the gamont surface. Midway in microgamete formation, the nucleus in the microgamete bud contains equal amounts of electron-lucent and electron-dense chromatin, and an apical cap consisting of two closely applied membranes forms over the anterior end. Mature microgametes separate from the gamont surface, leaving the residual body surrounded by a single membrane and containing numerous ribosomes, endoplasmic reticulum, and a few micronemes (Figure 1.11). Microgametes are rod shaped (1.4 × 0.5 µm for C. parvum), with a flattened anterior end. They lack both flagellae and mitochondria typically observed in microgametes of other coccidia. Most of the
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FIGURE 1.16 TEMs of C. parvum oocyst walls. (A) Tangential section through wall indicates that the outer electrondense layer has a network structure and the inner layer has a linearity suggestive of a filamentous array (arrowheads) (Magnification ×14,000). Harris and Petry, 1999. J. Parasitol. 85, 839–849. With Permission). (B) Excysted oocyst containing residuum, walls parted and curled inward at the suture opening (arrowheads) (Magnification ×14,000).
microgamete consists of a condensed nucleus (Figure 1.12). A plasmalemma completely surrounds the body. Beneath it, a single membrane extends posteriorly approximately two-thirds the body length. Originating at an anterior conical structure, 8 microtubules extend posteriad in close proximity to the surface of the nucleus (Figure 1.12). Three to five concentric lamellae extend outward at 90˚ to the long axis at the posterior margin of the apical cap (Figure 1.12), their function unknown. Electron-dense granules, also of undetermined function, are found in the cytoplasm at mid body (Figure 1.12). Macrogamonts of C. parvum are approximately 4 to 6 µm, spherical to ovoid, have a large central nucleus with a prominent nucleolus, and contain lipid bodies, amylopectin granules, and unique wallforming bodies in the cytoplasm (Figure 1.13). Little of the fertilization process of a macrogamont by a microgamete has been recorded, suggesting that the process is rapid. Microgametes attach at their apical cap to the surface of host cells harboring macrogamonts (Figure 1.12), pass through the surrounding membranes, and enter the cytoplasm of the macrogamont (Figure 1.13). Fusion of nuclei has not been observed. The fertilized macrogamont, or zygote, develops into an oocyst with either a thin or a thick wall (Figure 1.14). Those that develop into thick-walled oocysts have Type I and II wall-forming bodies similar to other coccidia (Figure 1.15). Those that develop into thin-walled oocysts lack the characteristic wall-forming bodies. Initially, two unit membranes form simultaneously external to the plasmalemma, whereas the sporont separates from the feeder/attachment organelle. Then, wall-forming body material is transported or exocytosed across the oocyst pellicle (i.e., plasmalemma and inner membrane) where it forms a thin, moderately coarse outer layer and a finely granular inner layer (Figures 1.15 and 1.16).
VII. Transmission Cryptosporidiosis is transmitted via the fecal-oral route by the oocyst stage. Sources and rates of infection are discussed in detail in Chapter 4. Cryptosporidium parvum is not only zoonotic but has been found to infect many species of mammals (Chapters 5, 16–18). Opportunities for infection of humans increase with close human-to-human contact and during care of infected livestock, zoo animals, or companion animals. Drinking water and recreational waters serve as vehicles for transmission, and many outbreaks have been reported from both sources (Chapters 11 and 12). Oocysts have been detected on fresh
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FIGURE 1.17 SEM of stages covering the villar and intervillar surface of the small intestine of an experimentally infected mouse (Magnification ×225). (From Vitovec, J. and B. Koudela, 1988. J. Vet. Med. 35, 515. With permission.)
FIGURE 1.18 SEM of the cloaca of a chicken experimentally infected with C. baileyi. Some villi are fused near the surface of the parasites. The host cell membrane covering one schizont is gone, exposing merozoites. Note the craters that remain after parasites have ruptured host cells at the attachment site. (Courtesy of W. L. Current and S. L. White, Lilly Research Laboratories, Indianapolis, IN.) (From Fayer, R. 1997. Cryptosporidium and Cryptosporidiosis. CRC Press, Boca Raton, FL. With permission.)
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vegetables and in irrigation waters, and linked to outbreaks of foodborne infection (Chapters 10 and 13). A “Fault Tree” indicating known and potential sources of infection is presented as Figure 1.1.
VIII. Prevention, Control, and Treatment As the sole mechanism for transmission, oocysts have evolved to be widely dispersed and to survive in harsh environments for long periods of time. They are highly resistant to natural stresses and many manmade chemical disinfectants. Oocysts probably evolved in dispersed mobile populations where there was strong selective pressure for long-term survival, the result being production of massive numbers of oocysts that survived for long periods of time (Blewett, 1989). In contrast, modern times are characterized by concentrated fixed populations of animals and people. The consequence is exposure of both populations to extremely high levels of infective organisms. To control infections in animal populations, one strategy might be to modify this imbalance by continuously moving animals to clean areas. However, where large numbers of domesticated animals are concerned, this is rarely economically possible. For human populations, disinfection procedures are sought to minimize person-to-person transmission in domestic and institutional settings and to deal effectively with contamination of recreational and drinking water. Because all infections with Cryptosporidium are initiated by ingestion or inhalation of the oocyst stage, measures to prevent or limit the spread of infection must be targeted to eliminate or reduce contamination from infectious oocysts in the environment. There are no universally effective drugs for prophylaxis or therapy for humans or animals that will prevent or stop oocyst production by infected individuals (Chapter 9). Hygiene, including disinfection, remains the most effective management tool. To determine the effectiveness of any anti-oocyst activity, there must be an assay to determine viability and infectivity after treatment. The current principal methods include: (1) animal infectivity using a variety of animal models, especially rodents (Chapter 19); (2) in vitro cell culture, which utilizes select cell types to support development through a portion of the life cycle (Chapter 20); and (3) excystation alone or combined with dye techniques, which estimates viability but cannot determine infectivity (Chapter 6).
A.
Physical Factors That Reduce Oocyst Viability
Laboratory studies have attempted to determine the reduction in the number of viable or infectious oocysts after exposure to various physical stresses such as heat, cold, irradiation, pressure, and desiccation (Table 1.7). At 5–15˚C, some oocysts of C. parvum, cleaned of feces and stored in water, remained infectious for 6 months although the number of infectious oocysts within the population slowly and steadily decreased with time of storage (Fayer et al., 1998). As temperatures decreased below 5˚C or increased above 15˚C, the survival time shortened (Table 1.8). The longevity of C. parvum oocysts at various temperatures appeared linked to carbohydrate energy reserves stored in sporozoites, and the residual bodies as amylopectin granules were consumed more rapidly in relationship to higher temperatures (Fayer et al., 1998; Jenkins et al., 2003). As demonstrated for other coccidian parasites, amylopectin provides the energy needed for excystation and host-cell invasion (Vetterling and Doran, 1969). Loss of infectivity at higher temperatures was also associated with decreased ATP content resulting from higher oocyst metabolic activity (King et al., 2005). Temperatures at or above 64.2˚C for 5 min and 72.4˚C for 1 min rendered oocysts noninfectious (Fayer, 1994). Oocysts of C. parvum suspended in water and whole milk were rendered noninfectious for mice when exposed to 71.7˚C for 5, 10, and 15 s in a laboratory pasteurizer (Harp et al., 1996). Oocysts of C. parvum withstood freezing at or above –20˚C for extended periods but did not survive –70˚C or lower even in the presence of cryoprotectants (Fayer and Nerad, 1996; Fayer et al., 1991). The effect of repeated freezing and thawing at –10˚C on oocysts in soil with 3 to 78% water content did not significantly differ from oocysts held at –10˚C under static conditions (Kato et al., 2002).
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TABLE 1.7 Physical Disinfection of Cryptosporidium parvum Oocysts Agent Heat Heat Heat Heat
Heat Heat (under pressure) Heat Freezing Freezing
Freezing Freezing
Water potential versus temperature Microwave Microwave e-beam irradiation Gamma irradiation Gamma irradiation Ultrasound
Conditions
Results NI NI NI Protein changes I NI I NI NI > 3 log reduction
In vivo In vivo In vivo DEP In vivo In vivo In vivo In vivo In vivo Cell culture
37°C, 48 h, in yogurt
20% reduction
Dyes
–196°C, 10 min –20°C, 3 days –70°C, 1 h –20°C, 8 h; 1 day –15°C, 24 h; 1 week –10°C, 1 week –20°C, 24 h ice cream, –20°C, 24 h water Multiple freeze–thaw cycles at –10°C in soil with varied moisture –4, –12, and –33 bars versus temperatures
NI NI NI I; NI I; NI I 100% reduction 92% reduction 99% reduction
In vivo In vivo In vivo In vivo In vivo In vivo Dyes Dyes Dyes
Degradation enhanced with stress
Dyes
Walker et al., 2001
0–45 s at 100% power
Inactivation was temperature dependent Slight reduction 100% reduction at 2.0 kGy
Dyes In vivo In vivo In vivo
Ortega and Liao, 2006 Collins et al., 2005a Collins et al., 2005a
100% reduction at 250 and 300 Gy 100% reduction at 450–500 Gy
In vivo
Jenkins et al., 1998a
In vivo
Jenkins et al., 2004
Possible reduction
Dyes In vivo In vivo In vivo In vivo Cell culture
Oyane et al., 2005
Zimmer et al., 2003
In vivo
Craik et al., 2001
Cell culture In vivo Cell culture In vivo Cell culture In vivo
Shin et al., 2001
1–3 s in oyster tissue 1.0, 1.5, and 2.0 kGy in oyster tissue 150, 200, 250, 300 Gy 350–500 Gy
UV
1 log reduction 2 log reduction 4 log reduction 1.5–3.2 log reduction 3.3–3.6 log reduction 2 and 3 log reduction at < 10 and 25 mJ/cm2 ~3 log reduction
UV UV UV UV
3 mJ/cm2 (MP) 14.3 mJ/cm2 in cider 20 mJ/cm2 (LP) 60 mJ/cm2 (MP)
90% reductiona 100% reduction > 2 log reduction 4.0–4.5 log reduction
UV
80 mJ/cm2
UV
120 mJ/cm2 (LP)
90% reduction 99% reduction 100% reduction
UV UV
Ref.
45°C, 20 min 60°C, 6 min 50–55°C, 5 min 121°C, 10 min 59.7°C, 5 min 64.2°C, 5 min 67.5°C, 1 min 72.4°C, 1 min 71.7°C, 5, 10, 15 s 121°C, 18 min
26.6 kHz, 30 W at 33 mL/min 0.48 mJ/cm2 0.97 mJ/cm2 1.92 mJ/cm2 1 to 3 mJ/cm2 (LP) 1.3 to 4 mJ/cm2 (MP) 0.8–119 mJ/cm2 (LP and MP) 3 mJ/cm2 (LP)
UV
Test
Anderson, 1985 Blewett, 1989 Archer et al., 1993 Fayer, 1994
Harp et al., 1996 Barbee et al., 1999 Deng and Cliver, 1999 Sherwood et al., 1982 Fayer and Nerad, 1996
Deng and Cliver, 1999 Kato et al., 2002
Morita et al., 2002
Excyst
Johnson et al., 2005 Hanes et al., 2002 Keegan et al., 2003 Belosevic et al., 2001 Ransome et al., 1993
In vivo
Drescher et al., 2001
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TABLE 1.7 (CONTINUED) Physical Disinfection of Cryptosporidium parvum Oocysts Agent
Conditions
Results
2
UV
8748 mJ/cm
Pulsed light Solar irradiation
1 J/cm2 830 W m2, 40°C, 10 h (30 kJ) 305, 370, 400, 480, and 550 MPa in oysters Air dried, 2 h Air dried, 4 h Air dried in feces, 1–4 days 10 min 1h 2h 4h
High pressure Drying Drying Drying
Test
100% reduction 100% reduction NI
Excyst Dyes In vivo In vivo
93.3% reduction at 550 MPa
In vivo
97% reduction 100% reduction NI 19% reduction 31% reduction 55% reduction 95% reduction
Excyst Dyes In vivo Dyes
Ref. Campbell et al., 1995 Dunn et al., 1995 McGuigan et al., 2006 Collins et al., 2005 Robertson et al., 1992 Anderson, 1986 Deng and Cliver, 1999
Note: I = infectious; NI = noninfectious; in vivo testing performed in mice; DEP = dielectrophoresis; Ex = excystation; a = C. hominis; dyes = usually DAPI and/or propidium iodide.
TABLE 1.8 Effect of Temperature Versus Storage Time on Infectivity of C. parvum Oocysts for Mice ˚C –10 –5 0 5 10 15 20 25 30 35 a
Number of Weeks Oocysts Stored before Bioassay 1 2 4 8 12 16 20 a
0 100 100 100 100 100 100 100 100 20
0 100 100 100 100 100 100 100 100 0
100 100 100 100 100 100 100 10
0 100 100 100 100 100 100 20
0 100 100 100 100 90 10 10
0 100 100 100 100 100 0 0
100 100 100 100 0 0
24
0 100 100 100 10 0
Percent of infected mice based on 7 to 10 mice examined per bioassay period.
Source: Adapted from Fayer, R., Trout, J.M. and Jenkins, M.C. 1998. Infectivity of Cryptosporidium parvum oocysts stored in water at environmental temperatures. J. Parasitol. 84, 1165–1169. With permission.
Several studies have shown the efficacy of ultraviolet (UV) irradiation for rendering oocysts noninfectious (Table 1.7). Although some investigators report variations in sensitivity of oocysts among batches tested (Craik et al., 2001; Sivaganesan and Sivaganesan, 2005), and levels of inactivation vary among studies, doses at or below 10 mJ/cm2 result in considerable reduction in oocyst infectivity (Table 1.7). One potential use of UV is for treatment of drinking water (Chapter 11). Of two UV sources, lowpressure (LP) and medium-pressure (MP) mercury lamps, the former has been used more extensively. LP sources emit a wavelength of ~254 nm, which closely approximates the maximum DNA absorption wavelength of 260 nm and the wavelength imparting the maximum biocidal effect. Although one concerning the application of UV technology has been the potential ability for microorganisms to repair UV-induced DNA damage, in most studies no detectable evidence of repair was found after exposure to LP or MP lamp irradiation.
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Limited studies have determined the effects of drying, high-pressure, ultrasound, microwaves, e-beam irradiation, solar radiation, and gamma irradiation on C. parvum oocyst infectivity (Table 1.7). Highpressure, e-beam, and gamma irradiation may find application for the processing of certain foods. Drying has universal application indoors and outdoors.
B.
Chemicals That Reduce Oocyst Viability
Numerous chemical disinfectants, most of which have been used for disinfection of bacteria and viruses, have been tested for efficacy against Cryptosporidium oocysts (Tables 1.9 to 1.11). Care must be taken in the interpretation of all test results associated with disinfection studies. Neither the incorporation of dyes nor the observation of excystation can validate either infectivity or the ability to complete the life cycle. Even the most realistic tests, employing animal models, especially rodents, must be interpreted with care (Chapter 19). However, in vivo testing has been conducted in only a minority of studies, perhaps because it is both expensive and time consuming. Some in vivo tests employing cell cultures provide a good alternative. TABLE 1.9 Commercial Chemical Disinfectants Tested Against Cryptosporidium Oocysts Disinfectant
Conditions
Result
Test
Reference
Tegodor (aldehydes) Formula H (aldehydes) Dikonit (chlorinated cyanuric acid) Jodonal (iodonal) Lastanox (tributyltinoxide) Mycolastanox (tributyltinoxide) Vincoa (formaldehyde based)
3%, 6 h 17%, 6 h 3% for 24 h 3% for 24 h 0.2% for 24 h 0.2% for 24 h 50%, 30 min, 25°C
I I I I I I NR
In vivo In vivo In vivo In vivo In vivo In vivo Excyst
Iofeca (iodine complex, phosphoric acid, phosphorous pentoxide) Pine Sola (pine oil, isopropyl alcohol) Creolina (coal oil, cresol)
50%, 30 min, 25°C
NR
Excyst
50%, 30 min, 25°C
NR
Excyst
50%, 30 min, 25°C
NR
Excyst
Microphenea (2-phenoxyethanol)
50%, 30 min, 25°C
NR
Excyst
DC&Ra (propanediol, ammonium chloride, formaldehyde) Nolvasana (chlorhexidine)
50%, 30 min, 25°C
NR
Excyst
50%, 30 min, 25°C
NR
Excyst
30 min @ 22°C 30 min @ 22°C 30 min @ 22°C Conc. not given 30 min, 22°C 5%, 30 min, 22°C 30 min @ 22°C
NR NR NR 95% R
Excyst Excyst Excyst Excyst
Angus et al., 1982 Angus et al., 1982 Pavlasek, 1984 Pavlasek, 1984 Pavlasek, 1984 Pavlasek, 1984 Sundermann et al., 1987 Sundermann et al., 1987 Sundermann et al., 1987 Sundermann et al., 1987 Sundermann et al., 1987 Sundermann et al., 1987 Sundermann et al., 1987 Blewett, 1989 Blewett, 1989 Blewett, 1989 Blewett, 1989
97% R NR
Excyst Excyst
Blewett, 1989 Blewett, 1989
1%, 30 min, 22°C 1%, 30 min, 37°C 30 min @ 22°C
20% R 90% R NR
Excyst
Blewett, 1989
Excyst
Blewett, 1989
30 min @ 22°C 30 min @ 22°C 30 min @ 22°C 4% 6%, 1 h, 37°C 2%, 1 h, 37°C
NR NR NR NR NR NR
Excyst Excyst Excyst Excyst Excyst Excyst
Blewett, 1989 Blewett, 1989 Blewett, 1989 Blewett, 1989 Holton et al., 1994 Holton et al., 1994
Buraton (ethanol) Microgen (paraformaldehyde) Dettol (parachlorometaxylenol) Exspor (chlorine dioxide) Oocide (ammonia) Savlon (chlorhexidine, centrimide, alcohol) FAM 30 (iodophore) Presept (sodium dichloroisocyanurate) Hycolin (synthetic phenol) Sporocidin (glutaraldehyde) Vircon (peroxygen) Lysol (ortho-dichloro-para-phenol) Sactimed (quaternary ammonia) Cidex (glutaraldehyde)
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Cryptosporidium and Cryptosporidiosis, Second Edition
TABLE 1.9 (CONTINUED) Commercial Chemical Disinfectants Tested Against Cryptosporidium Oocysts Disinfectant Cidex (glutaraldehyde) Virkon (peroxygen) Phoraid (iodophore) Pentapon HDY (beta-ene) Pentapon DC1 (beta-ene) Esculase (detergent) + Sporadyn (1.6 % glutaraldehyde) Esculase (detergent) + Cidex (2% glutaraldehyde) Esculase (detergent) + Hibitane (chlorhexidine) Product A (providone-iodine based)
Conditions
Result
Test
Reference Barbee et al., 1999 Holton et al., 1994 Holton et al., 1994 Holton et al., 1994 Holton et al., 1994 Vassal et al., 1998
Barbee et al., 1999 Barbee et al., 1999 Barbee et al., 1999
2.4%, 25°C, 45 min 1%, 1 h, 37°C 1%, 1 h, 37°C 5%, 1 h, 37°C 5%, 1 h, 37°C ~20°C, 10 min
0.3 log R NR NR NR SR NR
~20°C, 10 min
NR
~20°C, 10 min
NR
10%, 10–600 min, ~20°C
> 90% R excyst @ 600 min
Product B (phenol based)
10%, 10–600 min, 20°C
Minimal reduction
Product C (glutaraldehyde based)
2.5%, 10–600 min, 25°C
ND @ 600 min
Betadine (iodophore) Sporox (hydrogen peroxide) Sterrad 100 system (hydrogen peroxide) TBQ (quaternary ammonium) Vesphene IIse (phenolic) CIDEX-OPA (orthophthalaldehyde) Clinicide (detergent, quaternary ammonium) Omega (ammonium compounds) Wescodyne (iodine) Ox-Agua (hydrogen peroxide, silver nitrate) Ox-Virin (hydrogen peroxide, peracetic acid) Agri’germ 1000 (formaldehyde, ammonia, glutaraldehyde) Agroxyde II (Peracetic acid, hydrogen peroxide, acetic acid)
1% iodine, 20 min, 20°C 7.5%, 20 min, 20°C 6 mg/L
NR > 3 log R 3 log R
Cell culture Excyst Excyst Excyst Excyst In vivo (rats) In vivo (rats) In vivo (rats) Excyst and Cell culture Excyst and Cell culture Excyst and Cell culture Cell culture Cell culture Cell culture
1,128, 10 min, 20°C 1,128, 20 min, 20°C Undiluted, 20 min, 20°C
NR < 1 log R NR
Cell culture Cell culture Cell culture
Barbee et al., 1999 Barbee et al., 1999 Barbee et al., 1999
4, 13, 33 min @ 22°C
NR
Cell culture
Weir et al., 2002
4, 13, 33 min @ 22°C 4, 13, 33 min @ 22°C 10% for 1 h
NR NR I
Cell culture Cell culture In vivo
Weir et al., 2002 Weir et al., 2002 Quilez et al., 2005
3% for 30 min
I
In vivo
Quilez et al., 2005
30% for 1 h
5/23 mice
In vivo
10% for 1 h
3/28 mice
In vivo
Castro-Hermida et al., 2006 Castro-Hermida et al., 2006
Note:
Vassal et al., 1998 Vassal et al., 1998 Wilson and Margolin, 1999 Wilson and Margolin, 1999 Wilson and Margolin, 1999
a = Tested against C. baileyi oocysts; NR = no marked reduction; R = reduction; I = infection, ND = no development; SR = significant reduction.
Oocysts of C. parvum have shown considerable resistance against the effects of most commercial disinfectants, and the effective concentrations for others are generally not practical for disinfection outside the laboratory. The high concentrations required to significantly reduce oocyst infectivity are either very expensive, highly toxic, require an unacceptably long exposure time, or appear to be effective only at elevated temperatures. Among the commercial disinfectants that appear most effective are those that contain hydrogen peroxide, chlorine dioxide, or ammonia. Although bromine-, chlorine-, and iodine-related compounds can greatly reduce the ability of oocysts to excyst or infect, relatively high concentrations or long exposure periods are required, limiting most practical applications (Table 1.10). For applications to drinking and recreational water, see Chapters 11 and 12.
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TABLE 1.10 Halogen Disinfectants Tested Against Cryptosporidium Oocysts Disinfectants Bromine Chlorine
Chlorine Chlorine Chlorine Chlorine after shaking with sand Hypochlorite Hypochlorite Hypochlorite Hypochlorite Hypochlorite Hypochlorite Chlorine dioxide Chlorine dioxide Chlorine dioxide Chlorine dioxide Exspor (Chlorine dioxide) Chloramine Monochloramine Monochloramine Monochloramine Iodophore Provid (Iodine) Iodine Iodine bromide
Conditions
Results
Test
Reference
1180 mg/L, 60 min 5 mg/L 80 mg/L, 2 h 80 mg/L, 2 h 867–5118 mg/L, 24 h 16000 mg/L, 12 h 28000 mg/L, 24 h 80 ppm, 90 min 1 mg/L, 5 min, 25˚C
89% reduction nr 100% reduction 99% reduction 73% to 88% reduction 90% reduction 100% reduction 90% reduction reduction
Excyst Excyst Excyst In vivo Excyst Excyst
Ransome et al., 1993 Quinn and Betts, 1993
Ransome et al., 1993 Smith et al., 1990
Excyst Excyst
Korich et al., 1990 Parker and Smith, 1993
2.8%, 30 min, 25˚C 1%, 30 min, 22˚C 1%, 30 min, 37˚C 3%, 18 h 5.25%, 2 h, 20˚C 5.25%, 10 min, 20˚C 6%, 33 min 0.07 mg/L, 16 min 0.22 mg/L, 30 min 0.6 mg/l, 1 h 0.6 mg/l, 90 min 1.3 mg/L, 1 h
89% reduction 55% reduction 69% reduction I I < 1 log reduction NR 97% reduction 94% reduction NR 90% reduction 93% reduction
Excyst Excyst
Sundermann et al., 1987 Blewett, 1989
In vivo In vivo Cell culture Cell culture Excyst In vivo Excyst In vivo Excyst
Campbell et al., 1982 Fayer, 1995 Barbee et al., 1999 Weir et al., 2002 Peeters et al., 1989
4.03 mg/L, 15 min
96% reduction
Excyst
Ransome et al., 1993
Conc. not given 30 min, 22˚C 3%, 24 h 80 ppm, 90 min 0.066 mg/L, 48 h 3.76 mg/L, 24 h 80 mg/L, 2 h 80 mg/L, 90 min 4%, 18 h 10%, 30 min, 22˚C 10%, 30 min, 37˚C 10 mg/L, pH4-7, 1h 39.1 mg/L, 60 min
95% reduction
Excyst
Blewett, 1989
I 90% reduction 81% reduction 77% reduction 100% reduction 90% reduction I 19% reduction 53% reduction 85–56% reduction
In vivo Excyst Excyst
Pavlasek, 1984 Korich et al., 1990 Ransome et al., 1993
Excyst In vivo In vivo Excyst
Ransome et al., 1993 Campbell et al., 1982 Blewett, 1989
Excyst
Ransome et al., 1993
72% reduction
Excyst
Ransome et al., 1993
Peeters et al., 1989 Korich et al., 1990
Exposure to aqueous or gaseous ammonia and to hydrogen peroxide greatly reduced or eliminated oocyst infectivity, suggesting applicability (Table 1.11). Ozone appeared to be one of the most effective chemical disinfectants and has application against oocysts in water (Chapter 11).
C.
Environmental Factors That Potentially Reduce Oocyst Numbers
Potential beneficial effects in reducing the number of oocysts that enter surface waters have been associated with vegetative buffer strips (Atwill et al., 2002), constructed wetlands (Thurston et al., 2001; Nokes et al., 2003; Karim et al., 2004; Betancourt and Rose, 2005) and ponds (Grimason et al., 1993; Araki et al., 2001).
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TABLE 1.11 Miscellaneous Chemical Disinfectants Tested Against Cryptosporidium Oocysts Disinfectant Ethanol Ethanol Ethanol Propanol Isopropanol Isopropanol Methanol Glutaraldehyde Formaldehyde Formaldehyde Formaldehyde Formalin Formol saline Ammonia Ammonia Ammonia Ammonia Ammonia Parson’s ammonia Oocide (ammonia) Benzylkonium Campden solution Carbon monoxide Cresylic acid Bromomethane Ethylene oxide Ethylene oxide Hydrogen peroxide Hydrogen peroxide Hydrogen peroxide Hydrogen peroxide (gas plasma) Hydrogen peroxide
Peracetic acid Peracetic acid (Aldrich) Peracetic acid (Steris) Phenol Potassium dichromate Potassium permanganate
Conditions
Results
90%, 30 min, 22 and 37°C 70%, 20 min, 20°C 70%, 33 min, 24°C 90%, 30 min, 22 and 37°C 90%, 30 min, 22 and 37°C 70%, 33 min, 24°C 37%, 33 min, 24°C 2%, 30 min, 22°C 2%, 30 min, 37°C 1%, 30 min, 22°C 1%, 30 min, 37°C 5%, 24 h 100% gas, 24 h 10%, 1 and 7 days
NR NR NR NR NR NR NR NR 99% 36% 87% I NI 87%
10%, 18 h 1%, 30 min, 22°C 1%, 30 min, 37°C 5%, 18 h 100% gas, 24 h 150 mg/L 0.05 M, 24 h 50%, 30 min, 25°C 5%, 30 min, 22°C 10%, 18 h 0.5%, 30 min, 22°C 0.5%, 30 min, 37°C 100% gas, 24 h 5%, 18 h 100% gas, 24 h 100% gas, 24 h
NI > 60% reduction > 94% reduction NI NI 76–85% reduction 75% reduction 97% reduction 97% reduction I NR NR I I NI NI
450–500 mg/L @ 55–60°C 10 vol, 30 min, 22°C 306 mg/L, 10 min 288 mg/L, 30 min 6%, 10 min 6%, 20 min Low temperature 6%, 4 min 6%, 13 min 6%, 33 min 0.2%, 30 min, 22°C 0.2%, 30 min, 37°C 0.35%, 20 min, 20°C 0.2%, 12 min, 50°C 0.2%, 12 min, 25°C 0.2%, 12 min, 50°C 1%, 30 min, 22°C 1%, 30 min, 37°C 5%, 1 and 7 days
> 3 log reduction 95% reduction 95% reduction 98% reduction 550× reduction 1000× reduction Disinfection of endoscopes 1000× reduction NI NI 40% reduction 90% reduction NR 1.8 log reduction NR 1.8 log reduction NR NR 42% reduction
1%, 30 min, 22°C 1%, 30 min, 37°C
60% reduction > 95% reduction
reduction reduction reduction
reduction
Test
Reference
Excyst Cell culture Cell culture Excyst Excyst Cell culture Cell culture Excyst
Blewett, 1989 Barbee et al., 1999 Weir et al., 2002 Blewett, 1989 Blewett, 1989 Weir et al., 2002 Weir et al., 1999 Blewett, 1989
Excyst
Blewett, 1989
In vivo In vivo Excyst Dye In vivo Excyst
Pavlasek, 1984 Fayer et al., 1996 Campbell et al., 1993 Campbell et al., 1982 Blewett, 1989
In vivo Campbell et al., 1982 In vivo Fayer et al., 1996 Excyst Ransome et al., 1993 Excyst, Dye Jenkins et al., 1998 Excyst Sundermann et al., 1987 Excyst Blewett, 1989 In vivo Campbell et al., 1982 Excyst Blewett, 1989 In In In In
vivo vivo vivo vivo
Cell culture Excyst Excyst
Fayer et al., 1996 Campbell et al., 1982 Fayer et al., 1996 Fayer et al., 1996 Barbee et al., 1999 Blewett, 1989 Ransome et al., 1993 Barbee et al., 1999
ND
Vassal, et al., 2000
Cell culture
Weir et al., 2002
Excyst
Blewett, 1989
Cell culture
Barbee et al., 1999
Cell culture
Barbee et al., 1999
Excyst
Blewett, 1989
Excyst Dyes Excyst
Angus et al., 1982 Blewett, 1989
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TABLE 1.11 (CONTINUED) Miscellaneous Chemical Disinfectants Tested Against Cryptosporidium Oocysts Disinfectant Sodium hydroxide Sodium sulfite Sulfurous acid
Conditions 1%, 30 min, 22°C 1%, 30 min, 37°C 2–960 mg/L, 60 min 2–1000 mg/L, 60 min
Results > 75% reduction > 85% reduction NR NR
Test
Reference
Excyst
Blewett, 1989
Excyst Excyst
Ransome et al., 1993 Ransome et al., 1993
Note: NR = no marked reduction; I = infectious for mice; NI = not infectious for mice; ND = no data.
Under experimental conditions rotifers, which occupy niches in seawater, rivers, lakes, and ponds, and predaceous protozoa have been found to ingest oocysts of C. parvum (Fayer et al., 2000; Stott et al., 2003). Whether these predatory invertebrates digest and destroy the oocysts is not known. Shellfish including oysters, clams, mussels, and cockles are filter feeders that remove small particles from the surrounding aquatic environment. Oocysts have been detected in gills and the digestive diverticula of shellfish in numerous freshwater, and tidal and coastal water locations in North America and Europe (see review by Fayer et al., 2004). The overall impact on reduction of oocyst numbers, and whether oocysts lose infectivity after retention by shellfish, is unknown.
References Alvarez-Pellitero, P. and Sitja-Bobadilla, A. 2002. Cryptosporidium molnari n. sp. (Apicomplexa: Cryptosporidiidae) infecting two marine fish species, Sparus aurata L. and Dicentrarchus labrax L. Int. J. Parasitol. 32, 1007–1021. Alvarez-Pellitero, P., Quiroga, M.I., Sitja-Bobadilla, A., Redondo, M.J., Palenzuela, O., Padros, F., Vazquez, F. and Nieto, J.M. 2004. Cryptosporidium scophthalmi n. sp. (Apicomplexa, Cryptosporidiidae) from cultured turbot Scophthalmus maximus. Light and electron microscope description and histopathological study. Dis. Aquat. Organ. 62, 133, 2004. Anderson, D.R., Duszynski, D.W. and Marquardt, W.C. 1968. Three new coccidia (Protozoa, Telospora) from kingsnakes, Lampropeltis spp., in Illinois with a redescription of Eimeria zamenis Phisalix, 1921. J. Parasitol. 54, 577–581. Anderson, B. 1986. Effect of drying on the infectivity of cryptosporidia-laden calf feces for 3- to 7-day-old mice. Am. J. Vet. Res. 47, 2272 and 2273. Anderson, B.C. 1985. Moist heat inactivation of Cryptosporidium sp. Am. J. Pub. Hlth. 75, 1433–1434. Angus, K., Sherwood, D., Hutchison, G. and Campbell, I. 1982. Evaluation of the effect of two aldehyde-based disinfectants on the infectivity of faecal cryptosporidia for mice. Res. Vet. Sci. 33, 379–381. Anonymous. 1982. Cryptosporidiosis, assessment of chemotherapy of males with acquired immune deficiency syndrome (AIDS). Morbid. Mortal. Wkly. Rpt. 31, 89–102. Arcay, L., Baez de Borges, E. and Bruzual, E. 1995. Cryptosporidiosis experimental en la escala de vertebreados. I. Infecciones experimentales, II. Estudio histopatologico. Parasitol. al Dia 19, 20–29. Arcay de Peraza, L. and Bastardo de San Jose, T. 1969. Cryptosporidium ameivae sp. nov. Coccidia Cryptosporidiidae del intestino delgado de Ameiva ameiva de Venezuela. Acta Cient. Venezolona 20, 25. Archer, G.P., Betts, W.B. and Haigh, T. 1993. Rapid differentiation of untreated, autoclaved and ozone-treated Cryptosporidium parvum oocysts using dielectrophoresis. Microbios 73, 555–558. Araki, S., Martin-Gomez, S., Becares, E., Luis-Calabuig, E. and Rojo-Vasquez, F. 2001. Effect of high-rate algal ponds on viability of Cryptosporidium parvum oocysts. Appl. Environ. Microbiol. 67, 3322–3324. Asahi, H., Koyama, T., Arai, H., Kunakoshi, Y., Yamaura, H., Sirasaka, R. and Okutoni, K. 1991. Biological nature of Cryptosporidium sp. isolated from a cat. Parasitol. Res. 77, 237–240. Atwill, E.R., Hou, L., Karle, B.M., Harter, T., Tate, K.W. and Dahlgren, R.A. 2002. Transport of Cryptosporidium parvum oocysts through vegetated buffer strips and estimated filtration efficiency. Appl. Environ. Microbiol. 68, 5517–5527.
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Sam-Yellowe, T.Y. 1996. Rhoptry organelles of the apicomplexa: Their role in host cell invasion and intracellular survival. Parasitol. Today 12, 308–316. Sherwood, D., Angus, K.W., Snodgrass, D.R. and Tzipori, S. 1982. Experimental cryptosporidiosis in laboratory mice. Infect. Immun. 38, 471–475. Shin, O.-A., Linden, K.G., Arrowood, M.J. and Sobsey, M. 2001. Pressure UV inactivation and DNA repair potential of Cryptosporidium parvum oocysts. Appl. Environ. Microbiol. 67, 3029–3032. Sivaganesan, M. and Sivaganesan, S. 2005. Effect of lot variability on ultraviolet radiation inactivation kinetics of Cryptosporidium parvum oocysts. Environ. Sci. Technol. 39, 4166–4171. Slapeta, J. 2006. Cryptosporidium species found in cattle, a proposal for a new species. Trends Parasitol. 22, 469–474. Slavin, D. 1955. Cryptosporidium meleagridis (sp. nov.). J. Comp. Pathol. 65, 262–270. Smith, H.V., Smith, A.L., Girdwood, R.W.A. and Carrington, E.G. 1990. The effect of free chlorine on the viability of Cryptosporidium sp. oocysts isolated from human faeces, in Cryptosporidium in Water Supplies, Badenoch, J., Ed., Her Majesty’s Stationary Office, London, 185–204. Spano, F., Putignani, L., Naitza, S., Puri, C., Wright, S. and Crisanti, A. 1998. Molecular cloning and expression analysis of a Cryptosporidium parvum gene encoding a new member of the thrombospondin family. Mol. Biochem. Parasitol. 92, 147–162. Stott, R., May, E., Ramirez, E. and Warren, A. 2003. Predation of Cryptosporidium oocysts by protozoa and rotifers, implications for water quality and public health. Wat. Sci. Tech. 47, 77–83. Sundermann, C.A., Lindsay, D.S. and Blagburn, B.L. 1987. Evaluation of disinfectants for ability to kill avian cryptosporidium oocysts. Comp. Anim. Pract. 36–39. Thurston, J.A., Gerba, C.P., Foster, K.E. and Karpiscak, M.M. 2001. Fate of indicator microorganisms, Giardia, and Cryptosporidium in subsurface flow constructed wetlands. Wat. Res. 35, 1547–1551. Triffit, M. 1925. Observations on two new species of coccidia parasitic in snakes. Protozoology 1, 19–26. Tyzzer, E.E. 1907. A sporozoan found in the peptic glands of the common mouse. Proc. Soc. Exp. Biol. Med. 5, 12–13. Tyzzer, E.E. 1910. An extracellular coccidium, Cryptosporidium muris (gen. et sp. nov.) of the gastric glands of the common mouse. J. Med. Res. 23, 487–511. Tyzzer, E.E. 1912. Cryptosporidium parvum (sp. nov.), a coccidium found in the small intestine of the common mouse. Arch. Protistenkd. 26, 394–412. Tyzzer, E.E. 1929. Coccidiosis in gallinaceous birds. Am. J. Hyg. 10, 269–383. Tzipori, S. 1983. Cryptosporidiosis in animals and humans. Microbiol. Rev. 47, 84–96. Umemiya, R., Fukuda, M., Fujisaki, K. and Matsui, T. 2005. Electron microscopic observation of the invasion process of Cryptosporidium parvum in severe combined immunodeficiency mice, J. Parasitol. 91, 1034–1039. Upton, S.J. 2000. Suborder Eimeriorina Leger, 1911, in An Illustrated Guide to the Protozoa, 2nd edition, Lee, J.J., Leedale, G.F. and Bradbury, P., Eds., Society of Protozoologists, 1, 318. Upton, S.J. and Current, W.L. 1985. The species of Cryptosporidium (Apicomplexa, Cryptosporidiidae) infecting mammals. J. Parasitol. 71, 625–629. Vassal, S., Favennec, L., Ballet, J.J. and Brasseur, P. 1998. Lack of activity of an association of detergent and germicidal agents on the infectivity of Cryptosporidium parvum oocysts. J. Infect. 36, 245–247. Vassal, S., Favennec, L., Ballet, J.J. and Brasseur, P. 2000. Disinfection of endoscopes contaminated with Cryptosporidium parvum oocysts (letter). J. Hosp. Infect. 44, 151. Vetterling, J.M. and Doran, D.J. 1969. Storage polysaccharide in coccidial sporozoites after excystation and penetration of cells. J. Protozool. 16, 772–775. Vetterling, J.M., Jervis, H.R., Merrill, T.G. and Sprinz, H. 1971. Cryptosporidium wrairi sp. n. from the guinea pig Cavia porcellus with an emendation of the genus. J. Protozool. 18, 243–247. Walker, M., Leddy, K. and Hagar, E. 2001. Effects of combined water potential and temperature stresses on Cryptosporidium parvum oocysts. Appl. Environ. Microbiol. 67, 5526–5529. Weir, S.C., Pokorny, N.J., Carreno, R.A., Trevors, J.T. and Lee, H. 2002. Efficacy of common laboratory disinfectants on the infectivity of Cryptosporidium parvum oocysts in cell culture. Appl. Environ. Microbiol. 68, 2576–2579. Weisburger, W.R., Hutcheon, D.F., Yardley, J.H., Roche, J.C., Hillis, W.D. and Charache, P. 1979. Cryptosporidiosis in an immunosuppressed renal-transplant recipient with IgA deficiency. Am. J. Clin. Pathol. 72, 473–478.
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Wetzel, D.M., Schmidt, J., Kuhlenschmidt, M.S., Dubey, J.P. and Sibley, L.D. 2005. Gliding motility leads to active cellular invasion by Cryptosporidium parvum sporozoites. Infect. Immun. 73, 5379–5387. Wetzel, R. 1938. Ein neus Coccid (Cryptosporidium vulpis ap. Nov.) aus dem Rotfuchs. Arch. wissenschaftl. prakt. Tierheilk. 74, 39 and 40. Wilson, J.A. and Margolin, A.B. 1999. The efficacy of three common hospital germicides to inactivate Cryptosporidium parvum oocysts. J. Hosp. Infect. 42, 231–237. Xiao, L., Bern, C., Limor, J., Sulaiman, I., Roberts, J., Checkley, W., Cabrera, L., Gilman, R.H. and Lal, A.A. 2001. Identification of 5 types of Cryptosporidium parasites in children in Lima, Peru. J. Infect. Dis. 183, 492–497. Zimmer, J.L., Slawson, R.M. and Huck, P.M. 2003. Inactivation and potential repair of Cryptosporidium parvum following low-pressure and medium-pressure ultraviolet irradiation. Wat. Res. 37, 3517–3523.
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2 Genomics
Jessica C. Kissinger
CONTENTS I. II.
Introduction .................................................................................................................................... 43 Molecular Data Types .................................................................................................................... 44 A. Karyotype .......................................................................................................................... 44 B. Expressed Sequence Tags ................................................................................................. 44 C. Genome Survey Sequences............................................................................................... 44 D. Genomic Sequences .......................................................................................................... 45 III. Genome Properties ......................................................................................................................... 45 A. Coding Capacity................................................................................................................ 46 B. Annotation ......................................................................................................................... 46 C. Functional Genomic Analyses .......................................................................................... 47 IV. Gene Regulation ............................................................................................................................. 48 V. Genome Resources......................................................................................................................... 48 VI. Comparative Genomics .................................................................................................................. 48 A. Phylogenetics..................................................................................................................... 49 B. Horizontal Gene Transfer.................................................................................................. 51 VII. Prospects......................................................................................................................................... 51 Acknowledgments.................................................................................................................................... 53 References............................................................................................................................. 53
I.
Introduction
In the 10 years that have elapsed since the first edition of Cryptosporidium and Cryptosporidiosis (Fayer, 1997), our knowledge of the molecular biology of Cryptosporidium has grown from a glimpse of a karyotype and a few individual gene sequences to two complete genome sequences, one complete chromosome 6 sequence, several molecular libraries, and databases that provide access to this growing body of data. Advances in Cryptosporidium culture, parasite isolation, and molecular extraction have provided the starting material, which, upon analysis, has generated significant insight into the basic biology, metabolism, and evolution of Cryptosporidium. These data have also provided the markers necessary to further refine identification approaches and characterize the structure and diversity of Cryptosporidium populations. This chapter will review the major molecular sequence resources that exist and highlight the insights that each data type has revealed. It begins with the Cryptosporidium parvum karyotype and proceeds through the various sequence data sets that have been generated, including the complete genome sequence, and ends with evolutionary insights derived from comparative genomics and future prospects.
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II. A.
Cryptosporidium and Cryptosporidiosis, Second Edition
Molecular Data Types Karyotype
Cryptosporidium parvum has a small genome for a eukaryote. Early characterization of the C. parvum karyotype revealed five relatively small (1 Mb or less) electrophoretic bands on a gel (Mead et al., 1988). Later, estimates of eight chromosomes appeared in the literature (Hays et al., 1995). With the aid of scanning densitometry, the genome size was estimated at 10.4 Mb (quite small for a eukaryotic organism), and eight chromosomes were estimated (Blunt et al., 1997). Soon thereafter, using DNA restriction analysis with rare cutting enzymes, it was discovered that several chromosomes of relatively similar size were comigrating on electrophoretic gels (hence the five electrophoretic bands observed earlier), and the authors confirmed a chromosome number of eight and genome size 10.4 Mb (Caccio et al., 1998; Putignani et al., 1999). Physical mapping of the genome by traditional genetic crosses is not possible for Cryptosporidium because of experimental limitations. However, a physical map containing hundreds of markers has been created for two isolates of C. parvum (Moredun and IOWA) via the HAPPY map technique (Piper et al., 1998; Bankier et al., 2003). The maps are collinear within expected margins of error. The map consists of 10 linkage groups that can be placed onto 8 chromosomes. These HAPPY maps were extremely useful in subsequent genome sequence efforts as independent verification for correct assembly of shotgun sequences and to provide a scaffold framework for the ordering and orientation of contigs generated by the genome sequence projects (Abrahamsen et al., 2004; Xu et al., 2004).
B.
Expressed Sequence Tags
There is only one published expressed sequence tag (EST) data set for C. parvum. It consists of 567 ESTs (~283,000 nt) generated from purified excysted sporozoites (Strong and Nelson, 2000). When clustered, these ESTs represent approximately 400 unique gene sequences. Analyses of the ESTs at the time of publication revealed that 32% were similar to existing sequences in the GenBank database; this number is higher today because of increased representation of the Apicomplexa within the database. When combined with 2000 of the GSS sequences discussed in the following text, a joint analysis by Strong and Nelson revealed a number of putative therapeutic targets including S-adenosylhomocysteine hydrolase, histone deacetylase, polyketide/fatty-acid synthases, cyclophilins, thrombospondin-related cysteine-rich protein, and ABC transporters. Due to the difficulty of purifying subsequent C. parvum developmental stages from host cells, there are no other published large-scale studies of Cryptosporidium gene expression. The 567 C. parvum ESTs stand in stark contrast to other apicomplexan species. There are 128K ESTs for Toxoplasma gondii, ~20K ESTs each for four human species of Plasmodium, and over 34K for Eimeria tenella (www.ncbi.nlm.nih.gov/dbEST/dbEST_summary.html).
C.
Genome Survey Sequences
There are 9145 genome survey sequences (GSSs) present in the NCBI GenBank as of December 2006. They represent three genome-level studies (Piper et al., 1998; Liu et al., 1999; Strong and Nelson, 2000) and a targeted high-density sequence of C. parvum IOWA chromosome VI (Piper et al., 1998; Bankier et al., 2003). These sequence analyses provided the first glimpse of C. parvum genome structure and a good estimate of the overall AT content of the genome at approximately 68%. In particular, the comprehensive analysis of chromosome 6 (1.16 Mb) revealed a dense genome, with tightly packed genes containing relatively small introns at a frequency of 25% (Table 2.1). Numerous homopolymer stretches of A’s and T’s ≥ 10 nucleotides in length were observed, as were two palindromic octamer motifs TGGCGCCA and TGCATGCA. An analysis of the predicted genes on the chromosome VI sequence revealed that C. parvum coding sequences contain relatively few introns and are longer than those for eukaryotes such as Saccharomyces or Caenorhabditis elegans, with an average of 616 amino acids, but
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TABLE 2.1 Comparison of Cryptosporidium Genome Annotation
Genome size Annotated genes Protein coding tRNA rRNA snoRNA Other* Hypothetical proteins Number of coding nucleotides Mean gene length Genes with introns Genes with introns, % of total Total number of introns Total intron sequence Average intron size
C. parvum Genome
C. hominis Genome
C. parvum Chr 6
9,087,724 (bp) 3,885 3,396 45 15 20 409 1,274 6,280,357 (bp) 1,806 (bp) 35 0.9 45 4,024 (bp) 89 (bp)
8,743,570 (bp) 3,956 3,886 46 24 0 0 2,377 5,304,055 (bp) 1,340 (bp) 7 0.17 12 637 (bp) 53 (bp)
1,330,257 (bp) 557 480 8 0 3 66 438 882,392 (bp) 1,797 (bp) 7 1.26 7 1,058 (bp) 151 (bp)
C. parvum Chr6–BX526834 1,164,853 (bp) 481 473 8 0 0 0 58 877,514 (bp) 1,824 (bp) 122 25.4 213 32,902 (bp) 154 (bp)
Note: Data calculated from Version 1.0 genome records present in GenBank in December 2006 (C. parvum AAEE01000000; C. hominis AAEL01000000; C. parvum chromosome 6 BX526834). Other* = identified coding regions without corresponding annotated protein sequences.
shorter than the average length of P. falciparum genes (700aa). Two additional features of the predicted gene set were unusual. First, 25% of the genes encode a transmembrane domain, suggesting a surface location; and second, only 43% of the predicted proteins contained identifiable motifs as compared to 55 ± 6% for other fully sequenced genomes available at the time (Bankier et al., 2003).
D.
Genomic Sequences
2004 was a pivotal year for sequence-based studies of Cryptosporidium. Genome sequences for two species, C. parvum IOWA type 2 and C. hominis TU502, were published (Abrahamsen et al., 2004; Xu et al., 2004). The genome sequences were generated via random shotgun sequencing approaches and yielded 13X and 12X coverage, respectively. Both genome assemblies were able to utilize the Cryptosporidium HAPPY map to provide a scaffold and order and orient the sequence contigs. The C. hominis genome sequence consists of 1422 contigs, and the C. parvum genome sequence, 18 contigs. Based on the genome sequences, the size of the C. parvum genome was determined to 9.11 Mb, slightly smaller than the estimated 10.4 Mb. The C. hominis genome sequence is 9.16 Mb. Most apicomplexan organisms examined to date contain three genome sequences: a nuclear genome, a mitochondrial genome, and an apicoplast genome. This is not the case with C. parvum, and most likely, all Cryptosporidium species. There is evidence of a relict mitochondrial organelle, but this relict organelle does not contain a genome sequence (Riordan et al., 2003; Putignani et al., 2004). In contrast to the relict mitochondrion, C. parvum does not contain an apicoplast organelle at all, and there is no trace of the apicoplast genome (Zhu et al., 2000b), making this organism quite distinct from other apicomplexan species examined thus far. As a result, all genome and gene content discussions will focus on the Cryptosporidium nuclear genome sequence.
III. Genome Properties In the interval that passed between the early Cryptosporidium EST and GSS studies, several other apicomplexan EST and genome projects were published: Toxoplasma gondii (Ajioka et al., 1998), P. falciparum (Gardner et al., 2002), P. yoelii (Carlton et al., 2002), Eimeria (Wan et al., 1999; Ng et al., 2002), Sarcocystis (Howe 2001), and others. The data from these sequence projects provided significantly
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enhanced resources for the identification of apicomplexan (Li et al., 2003a) and Cryptosporidium genes and motifs.
A.
Coding Capacity
Not unexpectedly, the nucleotide sequence and coding capacity of the C. hominis and C. parvum genomes are quite similar. They share an average nucleotide identity of 96.7% and a largely overlapping predicted complement of genes, 3956 in C. hominis and 3885 in C. parvum (Xu et al., 2004; Table 2.1). They have a nearly identical number of predicted tRNA and rRNA genes. Relative to other sequenced apicomplexan genomes, the Cryptosporidium genomes are reduced and contain ~1200 fewer genes than P. falciparum and ~3800 fewer than Toxoplasma gondii. Much, but not all, of the difference in gene number relative to P. falciparum can be attributed to a lack of nuclear-encoded genes targeted to the mitochondrion and apicoplast, and species-specific gene families of extracellular antigenic proteins (Gardner et al., 2002). Relative to other apicomplexan species and eukaryotes in general, Cryptosporidium has a highly reduced number of introns. The estimates in Cryptosporidium range from 5 to 20% (Bankier et al., 2003; Abrahamsen et al., 2004; Xu et al., 2004) for the number of genes containing introns in contrast to 54% in P. falciparum (Gardner et al., 2002) and 74% in Theileria parva (Gardner et al., 2005). Depending on the annotation, the number of hypothetical proteins ranges from 30 to 60% (Table 2.1). An analysis of the gene content for both species has revealed a number of missing biochemical pathways and a concomitant increase in the number of transporters. There is no evidence for a TCA cycle or an oxidative phosphorylation pathway. The pathways for the synthesis of many amino acids are missing. Conversely, the number of predicted transporters is at least double that observed in P. falciparum, with at least 12 sugar or nucleotide sugar transporters, 5 predicted amino-acid transporters, 3 fatty-acid transporters, and 23 ATP-binding cassette transporters. Although the mitochondrial genome is absent, there are several genes of mitochondrial origin in the nuclear genome. Expression of nuclear-encoded Cpn60 targets the mitochondria in Cryptosporidium (Riordan et al., 2003). Consistent with the lack of a functional mitochondrion and mitochondrial genome, Cryptosporidium pathways and gene content indicate an exclusively anaerobic lifestyle. A comprehensive overview of Cryptosporidium biochemistry is discussed in Chapter 3 and will be avoided here. There are many genes shared with other apicomplexan species for which we have genome data, but there are many differences as well. The conserved genes are primarily involved in core metabolism and DNA transcription and translation, and in components of apicomplexan-specific secretory organelles used during host-cell invasion. Differences include a lack of enzymes for de novo pyrimidine biosynthesis and the acquisition of nucleotide salvage enzymes. Cryptosporidium also has the capacity to synthesize polysaccharides, trehalose, and amylopectin. A number of genes of bacterial and even algal origin are present in the nuclear genome (Huang et al., 2004a; Madern et al., 2004; Striepen et al., 2004; Templeton et al., 2004a). A more detailed discussion of this class of genes appears in the following text. There are Cryptosporidium-specific gene families, such as the oocyst wall proteins (Xiao et al., 2000; Templeton et al., 2004b). However, no large arrays of variant surface antigens are present within the genome.
B.
Annotation
Annotation of the Cryptosporidium genomes has been difficult, and readers will undoubtedly notice that the numbers cited in this chapter do not necessarily reflect the published numbers or the statistics provided by various databases. This situation arises from the sheer lack of confirmatory experimental evidence to predict, verify, and correct gene predictions. As mentioned earlier, other apicomplexan genome projects have had the benefit of large numbers of ESTs and, in some cases, microarray and proteomics data to facilitate annotation. Cryptosporidium is not so fortunate. We are severely hampered by the lack of experimental data and comparative genomic data from several related Cryptosporidium species. As a result, the estimates of gene features that are hard to assign computationally, for example,
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introns, vary widely (Table 2.1). Adding to the difficulty is a lack of strain designation in some of the Cryptosporidium literature and, more recently, the worry that some strains might have diverged from original stocks (Cama et al., 2006). All these issues affect the ability of researchers and data curators to annotate the Cryptosporidium genome to the same level as other species. Hopefully, with the advent of improved Cryptosporidium culture and purification techniques, combined with advances in technology, additional Cryptosporidium genome sequences and new large-scale experimental data sets such as those described in the following text will emerge and facilitate additional annotation of Cryptosporidium genomes.
C.
Functional Genomic Analyses
Functional genomic analyses in Cryptosporidium are hampered by the availability of purified parasite material from intracellular developmental stages. These limitations do not apply to sporozoites and proteomic data that are available for sporozoites (Direct submission to CryptoDB.org from the Jonathan Wastling Laboratory) and from nonexcysted and excysting sporozoites (Snelling et al., 2006). These data can be used to enhance the existing annotation by providing evidence of expression for predicted and hypothetical proteins and, in some cases, proof of intron splicing. Snelling et al. (2006) were able to identify 303 proteins, 56 of which were annotated as hypothetical. The addition of expression information for these 56 genes proves that at least portions of the hypothetical predictions are real. Excitingly, the process of excystation produced a measurable increase in the abundance of 26 proteins, including ribosomes, metabolic enzymes, heat shock proteins, three apicomplexan-specific proteins, and five Cryptosporidium-specific proteins. As the authors suggest, the phylum- and species-specific genes may present targets for vaccines or chemotherapeutics aimed at interfering with parasite invasion (Snelling et al., 2006). NIH has funded projects for Cryptosporidium via proteomics and structural genomics resource center initiatives (Table 2.2), and several crystal structures for C. parvum proteins were recently published (Vedadi et al., 2007). Large-scale EST studies and microarrays are still hindered by the lack of pure parasite RNA from intracellular developmental stages. However, as a result of the genome sequence, brute-force efforts are possible. Gene-specific primers can be designed for all or a portion of the predicted genes and semiquantitative polymerase chain reaction (PCR) used to examine expression profiles during the 72-h window during which Cryptosporidium can be cultured in vitro. Small applications of this approach have already been published for the Cryptosporidium oocyst wall protein (COWP) genes (Templeton et al., 2004b), verification of expression of horizontally acquired genes (Huang et al., 2004a), and several groups of genes identified in searches for putative promoter elements in C. parvum (Mullapudi et al., 2007).
TABLE 2.2 Cryptosporidium or Apicomplexan-Specific “Omic” Resources Resource
Description/URL
CryptoDB C. hominis Genome C. muris Genome C. parvum Genome Proteomics Research Centers Structural Genomics Resource Center ApiDB Bioinformatics Resource Center
http://CryptoDB.org http://www.hominis.mic.vcu.edu/g_overview.html http://msc.tigr.org/c muris/index.shtml http://cryptogenome.umn.org/ http://www.proteomicresource.org/ http://sgc.utoronto.ca/SGCWebPages/sgc.php?sr=Cryptosporidium http://ApiDB.org
Reference Puiu et al., 2004 Xu et al., 2004 Abrahamsen et al., 2004 Vedadi et al., 2007 Aurrecoechea et al., 2007
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IV.
Cryptosporidium and Cryptosporidiosis, Second Edition
Gene Regulation
Early examination of C. parvum gene expression utilized differential display to compare transcript profiles of infected and uninfected cells. This approach led to the cloning of two C. parvum genes and examination of their developmental expression profiles via reverse transcriptase-polymerase chain reaction (RT-PCR) (Abrahamsen et al., 1996; Schroeder et al., 1998, 1999). Subsequent experiments examined the expression profiles of interesting genes identified in earlier EST and genome sequence survey (GSS) studies. Developmental regulation of a family of thrombospondin (TSP) type-1 domain proteins was examined by Deng et al. (2002), and expression of the family of oocyst wall proteins and COWPs was examined by Templeton et al. (2004b). Their analyses showed that these genes are expressed 48 to 72 h postinfection, consistent with the timing of oocyst formation. Analysis of the genome sequences revealed that basal transcriptional machinery is present, but there is a noticeable lack of identifiable eukaryotic transcription factors (Abrahamsen et al., 2004; Xu et al., 2004). All Apicomplexa examined thus far (Cryptosporidium, Plasmodium, Toxoplasma, and Theileria) appear to have reduced transcriptional machinery compared to other eukaryotes, and classic sequence motifs for transcription factor binding sites have not been identified. A study that used the C. parvum genome sequence to identify and analyze conserved, overrepresented sequence motifs located in the cis regions of genes found a number of novel motifs that correlated with patterns of gene expression as determined by semiquantitative real-time PCR. These findings are only correlations and proof of a functional role for the motifs in gene regulation, which awaits functional assays that are currently unavailable for Cryptosporidium (Mullapudi et al., 2007).
V.
Genome Resources
Data relevant to Cryptosporidium research are stored in a variety of database resources. General resources include the NCBI GenBank, EMBL and DDBJ family of sequence databases, the Protein Structure Database (PDB), and the Kyoto Encyclopedia of Genes and Genomes (KEGG) metabolic database. Each of these resources contains data relevant to Cryptosporidium research. The sequence databases contain genomic, EST, and GSS data. PDB houses protein crystal structure information and KEGG has metabolic pathway reconstructions for most completed genome sequences. Table 2.2 lists a number of Internet-accessible resources that provide Cryptosporidium sequence or functional genomic data. These resources include access to genomic sequences and, in many cases, tools to further analyze and query the data via keyword and BLAST searches. One database resource, CryptoDB, and its parent Bioinformatic Resource Center database, ApiDB, provide integrated access to all available Cryptosporidium sequence data and proteomic data and the tools needed to query across the data types. For example, it is possible to ask for all genes with a signal peptide and evidence of EST and proteomic expression. The results of this query can be further refined to list those genes that have, or do not have, orthologs in other apicomplexan species.
VI. Comparative Genomics Analyses of the Cryptosporidium genome in comparison with other Apicomplexa and eukaryotes in general have provided several insights. As already mentioned, a comparison of C. hominis with C. parvum revealed remarkably similar genome sequence, gene content, and gene organization, suggesting that differences in host specificity for these species are caused by subtle changes (Xu et al., 2004). Relative to most other apicomplexan species, the Cryptosporidium genome is highly streamlined, but this phenomenon has also been observed in the piroplasms, Theileria parva (Gardner et al. 2005) and T. annulata (Pain et al. 2005), and indicates independent large-scale gene losses.
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FIGURE 2.1 Distribution of ortholog clusters across a cladogram of presumed Alveolate relationships (Levine, 1988). The species are as indicated. The numbers in parentheses following the species names are the number of protein-encoding genes analyzed. The numbers above the branches indicate the number of orthologous clusters of genes shared by that node/branch and all of its descendants. For example, 820 orthologous clusters of genes are shared by all of the organisms, whereas 455 clusters are unique to P. falciparum. Numbers below the branches indicate orthologous clusters that are not detected and presumably lost. The exact numbers of orthologous clusters fluctuate depending on the version of the genome annotation used and the parameters used to define an ortholog within the program OrthoMCL (Li et al., 2003b), but the overall pattern is consistent.
A comparison of the number of orthologous gene clusters shared with other apicomplexans and the nearest free-living relatives for which we have genome sequence, the ciliates, illustrates how variable these organisms are with respect to their gene content (Figure 2.1). The species contained within the supergroup Alveolata share only 820 orthologous gene clusters in common, and the Apicomplexa are united by sharing an additional 279 gene clusters to the exclusion of the ciliates. Each major lineage within the Apicomplexa is characterized by a large number of unique gene clusters relative to the other genera and species compared. These numbers are artificially inflated because of paltry and biased genome sample coverage for this diverse phylum. However, in cases for which we have two species within the same genus, i.e., P. falciparum and P. vivax, we still see a large number of presumably species-specific genes. In the case of Plasmodium, the large families of variant surface antigens account for a large fraction of the differences; however, this is not the case for Cryptosporidium because these types of genes are not found in the genome. When one compares genomes at the level of protein domain architecture, it has been shown that C. parvum contains several unique domain architectures and some architectures that are shared with other apicomplexans (Figure 2.2). A comprehensive bioinformatics examination of surface proteins in C. parvum and P. falciparum yielded several significant findings (Templeton et al., 2004a). The authors identified 32 widely conserved surface domains distributed in 51 protein sequences. Ten of these domains (TSP-1, Sushi/CCP, Notch/Lin1, etc.) have previously been described only in animals, whereas others have been detected in prokaryotes. Consistent with other reports, this study confirms the mosaic nature of the apicomplexan genome and suggests that gene transfer has played a significant role in the evolution of this lineage.
A.
Phylogenetics
Although there is general agreement that Cryptosporidium is an apicomplexan (Levine 1970), its exact placement within the Apicomplexa has been elusive. On the basis of developmental and life-cycle characteristics, Cryptosporidium has traditionally been placed within the coccidia, in particular with the non-cyst-forming coccidian (see Chapter 1 for more details). However, analyses of sequence data have
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Cryptosporidium and Cryptosporidiosis, Second Edition
FIGURE 2.2 (A color version of this figure follows page 242.) Domain organizations of a representative set of surface proteins from Cryptosporidium parvum (top panel) and orthologs common to Plasmodium falciparum (bottom panel). All proteins shown here have a signal peptide sequence represented by a rectangular box at the beginning of the architectures. The domains are labeled as in Templeton et al., 2004a. (Reproduced with permission from Genome Research; Templeton, T.J., Iyer, L.M., Anantharaman, V., Enomoto, S., Abrahante, J.E., Subramanian, G.M., Hoffman, S.L., Abrahamsen, M.S., and Aravind, L. 2004a. Comparative analysis of apicomplexa and genomic diversity in eukaryotes. Genome Res. 14, 1686–1695.)
been more ambiguous, and depending on the molecules examined, its exact phylogenetic placement is quite variable. When molecular sequence analysis, primarily ribosomal sequence analysis, was first applied to isolates of Cryptosporidium from a variety of hosts, a number of different species were distinguished, and several were reclassified (Morgan et al., 1999a, 1999b; Xiao et al., 1999a, 1999b). Four main species emerged from the molecular classification: C. parvum, C. muris, C. baileyi, and C. serpentis. Later, a new species, specific to the infection of humans, C. hominis, was described (Morgan-Ryan et al., 2002; see Chapters 1 and 5 for more detail). As more sequence data emerged from Cryptosporidium, other apicomplexans, and organisms representative of the tree of life, it was possible to examine the molecular phylogenetic placement of Cryptosporidium. In a study of another apicomplexan parasite group, the gregarines, a molecular phylogenetic analysis or 18S ribosomal sequences revealed that Cryptosporidium actually forms a sister group to the gregarines and is not located within the coccidia (Carreno et al., 1999). A placement outside of the coccidia, most likely at the base of the Apicomplexa, was also supported by a
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subsequent analysis that included not only ribosomal but also several protein-encoding genes (Zhu et al., 2000a). However, it has also become clear that, depending on the gene used for the phylogenetic analysis, a variety of phylogenetic placements are observed. Some of these differences are attributable to known phylogenetic artifacts (Zhu et al., 2000a), others represent either the mosaic nature of the Cryptosporidium genome and the genes contained within it, and some are attributable to the fact that we have extremely limited sequence information for the vast diversity present within the Apicomplexa. Thus, depending on the species included in the analysis, the results are variable. So, what if many genes are used? A concatenated set of 20 genes was used to test for the monophyly of Plasmodium and Cryptosporidium relative to the rest of the tree of life, and monophyly was supported (Templeton et al., 2004a). However, we do not yet know the placement of C. parvum within the phylum. In a recent series of articles (Thompson et al., 2005; Barta and Thompson, 2006), a call for a reevaluation of the phylogenetic position of Cryptosporidium has been issued. Although it is argued that Cryptosporidium does indeed share many characteristics with the coccidia, including environmental niches, it is clear from molecular data, and now from the complete genome sequence, that Cryptosporidium is something else. As data from more genera within the Apicomplexa are gathered, the exact position should become clearer.
B.
Horizontal Gene Transfer
Horizontal gene transfer (HGT) is the acquisition of genetic material (genes or genome segments) from outside of the recipient cell. This is in contrast to intracellular gene transfer (IGT), in which genetic material is relocated within nuclear or organellar compartments in the same cell. Within the Apicomplexa, there is a significant amount of IGT that has been documented as transfers from the mitochondrion and apicoplast to the nuclear genome (Foth et al., 2003; Riordan et al., 2003; Huang et al., 2004b; Huang and Kissinger, 2006) as well as acquisition of plant-like genes, perhaps from the algal endosymbiont that gave rise to the apicoplast (Dzierszinski et al., 1999; Nagamune and Sibley, 2006). There are also numerous studies that have alluded to, or proven, instances of HGT to Cryptosporidium or its ancestor (Striepen et al., 2002, 2004; Huang et al., 2004a,b; Madern et al., 2004; Templeton et al., 2004a). In several cases, the consequences of these transfers have been investigated, and perhaps no metabolic pathways have been as significantly impacted as nucleotide biosynthesis and salvage. Investigations of Cryptosporidium nucleotide metabolism reveal a complete absence of de novo purine and pyrimidine biosynthesis. Instead, a number of salvage enzymes are utilized to meet the parasite’s nucleic acid requirements (Figure 2.3). Whereas this finding in and of itself sets Cryptosporidium apart from other apicomplexans, it was additionally discovered that three of the salvage enzymes were acquired by gene transfer (IMPDH, TK, UK/UPRT) and that two are uniquely present in Cryptosporidium (TK and UK/UPRT) (Striepen et al., 2002, 2004). These acquired genes are essential to the survival of the parasite and may represent therapeutic targets. Although the hosts of Cryptosporidium have IMPDH genes, they have a eukaryotic copy of the gene, not the prokaryotic version as in Cryptosporidium. Perhaps there is enough discrimination potential between the two genes to develop a chemotherapeutic (Striepen and Kissinger, 2004; Umejiego et al., 2004).
VII. Prospects These are exciting times for Cryptosporidium research. The insights gained from the first two genome sequences were significant, and a third more distantly related genome sequence, C. muris, is in progress (Table 2.2). Examination of the genome sequences has revealed much of the metabolic repertoire, or lack thereof, of the parasite. At the same time, vulnerabilities have been identified that can lead to urgently needed new therapeutics. Comparative genomics has revealed a highly streamlined, yet mosaic, genome with numerous genes acquired from bacteria. The C. hominis and C. parvum genomes are highly similar, and it appears that the differences in their host preferences are encoded in small, single- or multinucleotide polymorphisms and not as differences in gene content. Comparisons with other apicomplexan parasites and their
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FIGURE 2.3 The C. parvum nucleotide biosynthetic pathway is a phylogenetic mosaic. Major enzymes within this pathway are indicated as ovals. Enzymes with a similar evolutionary history are shaded with identical colors. The enzymes labeled 4 and 8 show a strong phylogenetic association with eubacteria, those labeled 6 and 12 show an association with plants. The remaining enzymes do not show indications of transfer. Analyses presented here are based on the C. parvum Type 2 IOWA genome. “T” = transporter. Two arrows indicate two or more enzymatic steps. Most nucleoside mono- and diphosphate kinase and phosphorylase steps have been omitted for simplicity. A complete set of these genes is present in the genome. (1) Adenosine transporter, (2) adenosine kinase, (3) adenosinemonophosphate deaminase, (4) inosine monophosphate dedydrogenase, (5) guanosine monophosphate synthase, (6) uridine kinase/uracil phosphoribosyltransferase, (7) uracil phosphoribosyltransferase, (8) thymidine kinase, (9) ribonucleotide diphosphate reductase, (10) cytosine triphosphate synthetase, (11) deoxycytosine monophosphate deaminase, (12) dihydrofolate reductase-thymidylate synthase. Details are as in Striepen et al., 2004 and Huang and Kissinger, 2006. (Reprinted with permission from Striepen, B., Pruijssers, A.P.J., Huang, J., Li, C., Gubbels, M.J., Umejiego, N.N., Hedstrom, L., and Kissinger, J.C. 2004. Gene transfer in the evolution of parasite nucleotide biosynthesis. Proc. Natl. Acad. Sci., USA 101, 3154–3159; Huang, J., and Kissinger, J. 2006. Horizontal and intracellular gene transfer in the Apicomplexa: The scope and functional consequences. in Genome Evolution in Eukaryotic Microbes. Katz, L. and Bhattacharya D., Eds., Oxford University Press.)
free-living relatives hold the promise of providing insight into the evolution of these parasites and adaptations to their hosts. Functional genomic technologies hold promise for studies of gene regulation, parasite development, and host–parasite interactions if new culture and parasite purification techniques are developed or bruteforce approaches such as real-time PCR are applied to genome-scale expression studies. The genome sequence and bioinformatics databases can facilitate greater interactions between the different Cryptosporidium research communities. As Cryptosporidium oocysts are surveyed and typed
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with molecular sequence markers, these data can be mapped to the genome sequences and used to identify alleles and single-nucleotide polymorphisms (SNPs) and also populations of parasites, effectively uniting the epidemiological and molecular research communities (Widmer et al., 2002).
Acknowledgments The author would like to thank Chih-Horng Kuo for sharing unpublished data used in the creation of Figure 2.1 and the apicomplexan and ciliate genome sequencing consortiums and databases for making data publicly available and accessible. This work would not have been possible without the collaborative support of Drs. Boris Striepen and Mitchell Abrahamsen.
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Morgan–Ryan, U.M., Fall, A., Ward, L.A., Hijjawi, N., Sulaiman, I., Fayer, R., Thompson, R.C., Olson, M., Lal, A., and Xiao, L. 2002. Cryptosporidium hominis n. sp. (Apicomplexa: Cryptosporidiidae) from Homo sapiens. J. Eukaryot. Microbiol. 49, 433–440. Mullapudi, N., Lancto, C.A., Abrahamsen, M.S., and Kissinger, J.C. 2007. Identification of putative cisregulatory elements in Cryptosporidium parvum by de novo pattern finding. BMC Genomics 8, 13. Nagamune, K. and Sibley, L.D. 2006. Comparative genomic and phylogenetic analyses of calcium ATPases and calcium-regulated proteins in the apicomplexa. Mol. Biol. Evol. 23, 1613–1627. Ng, S.T., Sanusi Jangi, M., Shirley, M.W., Tomley, F.M., and Wan, K.L. 2002. Comparative EST analyses provide insights into gene expression in two asexual developmental stages of Eimeria tenella. Exp. Parasitol. 101, 168–173. Pain, A., Renauld, H., Berriman, M., Murphy, L., Yeats, C.A., Weir, W., Kerhornou, A., Aslett, M., Bishop, R., Bouchier, C., Cochet, M., Coulson, R.M., Cronin, A., de Villiers, E.P., Fraser, A., Fosker, N., Gardner, M., Goble, A., Griffiths-Jones, S., Harris, D.E., Katzer, F., Larke, N., Lord, A., Maser, P., McKellar, S., Mooney, P., Morton, F., Nene, V., O’Neil, S., Price, C., Quail, M.A., Rabbinowitsch, E., Rawlings, N.D., Rutter, S., Saunders, D., Seeger, K., Shah, T., Squares, R., Squares, S., Tivey, A., Walker, A.R., Woodward, J., Dobbelaere, D.A., Langsley, G., Rajandream, M.A., McKeever, D., Shiels, B., Tait, A., Barrell, B. and Hall, N. 2005. Genome of the host-cell transforming parasite Theileria annulata compared with T. parva. Science 309, 131–133. Piper, M.B., Bankier, A.T., and Dear, P.H. 1998. A HAPPY map of Cryptosporidium parvum. Genome Res. 8, 1299–1307. Puiu, D., Enomoto, S., Buck, G.A., Abrahamsen, M., and Kissinger, J.C. 2004. CryptoDB: The Cryptosporidium genome resource. Nucleic Acids Res. 32, in press. Putignani, L., Sallicandro, P., Alano, P., Abrahamsen, M.S., Crisanti, A. and Spano, F. 1999. Chromosome mapping in Cryptosporidium parvum and establishment of a long-range restriction map for chromosome VI. FEMS Microbiol. Lett. 175, 231–238. Putignani, L., Tait, A., Smith, H.V., Horner, D., Tovar, J., Tetley, L., and Wastling, J.M. 2004. Characterization of a mitochondrion-like organelle in Cryptosporidium parvum. Parasitology 129, 1–18. Riordan, C.E., Ault, J.G., Langreth, S.G., and Keithly, J.S. 2003. Cryptosporidium parvum Cpn60 targets a relict organelle. Curr. Genet. 44, 138–147. Schroeder, A.A., Brown, A.M., and Abrahamsen, M.S. 1998. Identification and cloning of a developmentally regulated Cryptosporidium parvum gene by differential mRNA display PCR. Gene 216, 327–334. Schroeder, A.A., Lawrence, C.E., and Abrahamsen, M.S. 1999. Differential mRNA display cloning and characterization of a Cryptosporidium parvum gene expressed during intracellular development. J. Parasitol. 85, 213–220. Snelling, W.J., Lin, Q., Moore, J.E., Millar, B.C., Tosini, F., Pozio, E., Dooley, J.S., and Lowery, C.J. 2006. Proteomic analysis and protein expression during sporozoite excystation of Cryptosporidium parvum (coccidia, Apicomplexa). Mol. Cell Proteomics. Striepen, B., and Kissinger, J.C. 2004. Genomics meets transgenics in search of the elusive Cryptosporidium drug target. Trends Parasitol. 20, 355–358. Striepen, B., Pruijssers, A.P.J., Huang, J., Li, C., Gubbels, M.J., Umejiego, N.N., Hedstrom, L., and Kissinger, J.C. 2004. Gene transfer in the evolution of parasite nucleotide biosynthesis. Proc. Natl. Acad. Sci., USA 101, 3154–3159. Striepen, B., White, M.W., Li, C., Guerini, M.N., Malik, S.B., Logsdon, Jr. J.M., Liu, C., and Abrahamsen M.S. 2002. Genetic complementation in apicomplexan parasites. Proc. Natl. Acad. Sci. USA 99, 6304–6309. Strong, W.B. and Nelson, R.G. 2000. Preliminary profile of the Cryptosporidium parvum genome: An expressed sequence tag and genome survey sequence analysis. Mol. Biochem. Parasitol. 107, 1–32. Templeton, T.J., Iyer, L.M., Anantharaman, V., Enomoto, S., Abrahante, J.E., Subramanian, G.M., Hoffman, S.L., Abrahamsen, M.S., and Aravind, L. 2004a. Comparative analysis of apicomplexa and genomic diversity in eukaryotes. Genome Res. 14, 1686–1695. Templeton, T.J., Lancto, C.A., Vigdorovich, V., Liu, C., London, N.R., Hadsall, K.Z., and Abrahamsen, M.S. 2004b. The Cryptosporidium oocyst-wall protein is a member of a multigene family and has a homolog in Toxoplasma. Infect. Immun. 72, 980–987. Thompson, R.C., Olson, M.E., Zhu, G., Enomoto, S., Abrahamsen, M.S., and Hijjawi, N.S. 2005. Cryptosporidium and cryptosporidiosis. Adv. Parasitol. 59, 77–158.
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3 Biochemistry
Guan Zhu
CONTENTS I. Introduction .................................................................................................................................... 57 II. Highly Streamlined Metabolism .................................................................................................... 58 III. Energy and Carbohydrate Metabolism .......................................................................................... 59 IV. Nucleotide Metabolism .................................................................................................................. 62 V. Fatty Acid Metabolism................................................................................................................... 62 VI. Polyamine Metabolism................................................................................................................... 65 VII. Amino Acid Metabolism................................................................................................................ 65 VIII. Structural Proteins .......................................................................................................................... 66 IX. Membrane Proteins and Transporters ............................................................................................ 67 X. DNA and RNA Metabolism........................................................................................................... 68 XI. Potential Drug Targets.................................................................................................................... 69 XII. Some Technical Perspectives Related to Biochemical Studies ..................................................... 70 Acknowledgments.................................................................................................................................... 71 References............................................................................................................................. 71
I.
Introduction
Although Cryptosporidium has been known for over 100 years, what happens within the parasite cells has been a complete mystery for a very long time. In fact, much of the current knowledge regarding its biochemistry and metabolism has been generated in the last decade or so. Because of the technical difficulties in obtaining large amounts of pure parasite materials, only a very limited number of early studies addressed the biochemical features in the oocysts and free sporozoites (Denton et al., 1996; Entrala and Mascaro, 1997). Studies on the genetics of Cryptosporidium had a relatively late start in comparison to Plasmodium falciparum and Toxoplasma gondii. Many early gene discovery studies attempted to follow those conducted in P. falciparum and T. gondii by focusing on critical genes that might be served as drug targets. However, few were successful, largely because of the unexpected biochemical divergence between Cryptosporidium and these model apicomplexans. Indeed, the recently completely sequenced C. parvum genome has revealed that this parasite differs significantly from other apicomplexans in almost all important metabolic pathways (see the following sections for details) (Abrahamsen et al., 2004). This observation is further confirmed by the large-scale sequencing of C. hominis genome, which comprises an identical set of genes differing from those in C. parvum by less than 5% at the nucleotide level (Xu et al., 2004). Many conventional enzymes or drug targets are either absent in Cryptosporidium or highly divergent from those found in P. falciparum and T. gondii (Abrahamsen et al., 2004; Thompson et al., 2005), which explains the failure in early cloning of these “imaginary” targets. It also explains why so many compounds that are effective against other apicomplexans, including intestinal coccidia, showed little efficacy against C. parvum.
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The biochemical divergence of Cryptosporidium from other apicomplexans is also reflective of the evolutionary history of this genus. Although Cryptosporidium has been traditionally considered to be an intestinal coccidian (under the class Coccidia and order Eimeriida), recent molecular phylogenetic reconstructions have consistently placed this genus at the base of the phylum Apicomplexa (Zhu et al., 2000a), or even as a sister to the gregarines (a group of apicomplexan parasite of invertebrates) (Carreno et al., 1999), suggesting that the Cryptosporidium genus is likely an early emerging branch within the phylum. The genome sequencing projects on C. parvum and C. hominis serve as a milestone in the history of Cryptosporidium research (Abrahamsen et al., 2004; Xu et al., 2004). However, early expressed sequence tag (EST) and random genome sequence survey (GSS) projects also deserve credit as a first and critical step in understanding the metabolism in C. parvum at systematic levels (Abrahamsen, 1999; Liu et al., 1999; Strong and Nelson, 2000a, 2000b). Data obtained from these sequencing projects jump-started our overall understanding of the molecular biology, biochemistry, and evolution of Cryptosporidium. The gene discovery phase in Cryptosporidium research is largely over. In the postgenome era, we can now focus more on the study of functions and biological roles of parasite genes and their products. Although many Cryptosporidium species have been described in the literature, almost all biochemical data available today are obtained by studying C. parvum. No biochemical work has been performed directly on C. hominis, but its metabolism is essentially identical to C. parvum, based on their genome compositions (Xu et al., 2004). We also assume that the major metabolic pathways between C. parvum and other species (such as those from birds, reptiles, and fishes) are likely identical too, but experiments are needed to confirm this assumption. This chapter will focus on the major metabolic pathways that are important for parasite survival and development, many of which may also be explored as potential molecular targets for developing therapeutics for the control or treatment of Cryptosporidium infection in humans and animals.
II.
Highly Streamlined Metabolism
The recently completed genome sequencing project revealed that C. parvum possesses a compact, ~ 9.1megabase (Mb) genome distributed in eight chromosomes and predicted to encode more than 3800 genes (Abrahamsen et al., 2004). The genome size is much smaller than some other apicomplexans, including P. falciparum (~22.9 Mb) and T. gondii (80 Mb) (Gardner et al., 2002; Kim and Weiss, 2004). The C. parvum genome contains ~70% AT, a small number of introns (i.e., in ~5% genes), and small-sized intergenic regions (average 566 bp). The genome project, in which total DNA was isolated for sequencing by a random shotgun approach, also confirms that Cryptosporidium lacks mitochondrial and apicoplast genomes that are found in all other apicomplexans examined so far. The lack of apicoplast genome in C. parvum was proposed earlier based on experimental data (Zhu et al., 2000c). The genome project not only confirms this observation, but further reveals the absence of any apicoplast-associated pathways such as the Type II fatty acid synthesis and isoprenoid synthetic enzymes. Although C. parvum lacks a mitochondrial genome and genes encoding Krebs cycle, this parasite does possess some genes that encode mitochondrial proteins, including some elements of a protein import apparatus (e.g., TOM40 and TIM17), chaperons (HSP70 and HSP65), solute transporters, Fe-S cluster assembling proteins (e.g., nifS, nifU, frataxin, and ferredoxin), a cyanine-resistant alternative oxidase (AOX), and a pyridine nucleotide transhydrogenase, indicating the presence of a remnant mitochondrion in this parasite (LaGier et al., 2003; Riordan et al., 2003; Abrahamsen et al., 2004; Roberts et al., 2004; Slapeta and Keithly, 2004; Suzuki et al., 2004; Keithly et al., 2005). Also unexpected is the presence of only two headpieces of a bacterial-type ATP synthase (i.e., α- and β-subunits), but not any other subunits, for which their function and localization remain to be determined. Cryptosporidium appears to have lost most (if not all) de novo synthetic capacities, including those to make fatty acids, amino acids, and nucleosides. It also lacks the shikimate pathway that is present in other apicomplexans. These observations indicate that Cryptosporidium has adapted to an extreme parasitic lifestyle and solely relies on the host to provide its nutrients. Indeed, genome analysis has revealed an expanded set of transporters for scavenging nutrients, including those for sugars, amino
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acids, and nucleosides (Abrahamsen et al., 2004). Metabolism in this parasite is highly streamlined in comparison to that of Plasmodium or Toxoplasma. Figure 3.1 summarizes the major metabolic pathways we know today. More details about these pathways are discussed as follows. However, interested readers may also visit a number of Internet resources related to the Cryptosporidium genomes and metabolism. These include the following: 1. CryptoDB (http://CryptoDB.org), which contains the whole genome sequences and annotations for C. parvum and C. hominis, as well as EST and GSS sequences. Datasets can be downloaded, browsed, and analyzed with provided bioinformatics and data-mining tools. Information is also updated regularly (Heiges et al., 2006). 2. Kyoto Encyclopedia of Genes and Genomes (KEGG)’s pathway database (http://www.genome. ad.jp/kegg/pathway.html), which maintains a collection of manually drawn maps on almost all major metabolic pathways (based on genome annotations) of a large number of organisms, including C. parvum, C. hominis, and other apicomplexans. Enzyme sequences are annotated and cross-linked to a number of other sequence or enzyme databases. 3. The original C. parvum genome project site at University of Minnesota (http://134.84.110.219/), in which visitors can browse the genome and annotated metabolic and regulatory pathways. However, the site is not actively updated, because it was originally established to facilitate communication during the annotation of the project.
III. Energy and Carbohydrate Metabolism Because of the lack of a Krebs cycle, Cryptosporidium may rely solely on glycolysis as its energy source. It can utilize polysaccharides, including amylopectin, amylose, and hexoses such as glucose and fructose (Figure 3.1). All enzymes catalyzing reactions from hexose to pyruvate are present in the genome, in which hexokinase (HK) consumes one ATP in activating hexose, whereas phosphoglycerate kinase (PGK) and pyruvate kinase (PK) may each produce two ATPs when a single hexose is completely converted into two pyruvate molecules. Unlike humans or other typical aerobic organisms that use an ATPdependent phosphofructokinase (ATP-PFK), but similar to some other microanaerobic protists including Trichomonas and Giardia, Cryptosporidium uses a pyrophosphate-dependent PFK (PPi-PFK) to economize ATP consumption, for which the activity was previously detected in oocysts (Denton et al., 1996). Because Cryptosporidium does not have a Krebs cycle to completely oxidize carbohydrates, this parasite can generate only 3 net ATP molecules from a single hexose, which is much fewer than those generated by the aerobic pathway (up to 36 ATPs), but significantly higher than typical glycolysis that uses ATPPFK (only 2 net ATPs). Phosphoenolpyruvate (PEP) may be directly converted to pyruvate by PK, or indirectly via PEP carboxylase (PEPCL), malate dehydrogenase (MDH), and malic enzyme (ME). Although a weak activity of glycerol kinase (GK) catalyzing the formation of glycerol from glycerol3P was previously reported in oocyst extracts (Entrala and Mascaro, 1997), its gene ortholog cannot be found in either the C. parvum or C. hominis genome, suggesting that glycerol is not one of the end products for generating another ATP as seen in some other protists including trypanosomes (Kralova et al., 2000). However, Cryptosporidium is able to use glycerol-3P to synthesize phospholipid or other complex lipids. Pyruvate may be converted to acetyl-CoA by a bifunctional pyruvate, pyruvate:NADP+ oxidoreductase (PNO), which contains a pyruvate-ferredoxin oxidoreductase (PFO) domain and an NADPH-cytochrome P450 reductase domain (CPR) (Rotte et al., 2001). The architecture of PNO is unique to Cryptosporidium, as it is not present in any other apicomplexans examined so far, but found only in a distant free-living protist, Euglena gracilis (Rotte et al., 2001). Whereas Euglena PNO apparently contains a signal peptide sequence and is localized in the mitochondria, CpPNO is found in the cytosol (Ctrnacta et al., 2006). Acetyl-CoA can be converted by acetyl-CoA carboxylase (ACC) to malonyl-CoA, which serves as the building block in synthesizing fatty acids and polyketides. This parasite possesses only one cytosolic ACC, lacking the plastid ortholog found in Toxoplasma and Plasmodium (Jelenska et al., 2001, 2002;
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FIGURE 3.1
Cryptosporidium and Cryptosporidiosis, Second Edition
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Gornicki, 2003). At least two organic end products can be formed from acetyl-CoA, including acetate by an acetate-CoA ligase (AceCL, also referred to as acetyl-CoA synthetase), in which an extra ATP molecule can be generated from AMP and PPi, and ethanol, by a bifunctional type E alcohol dehydrogenase (adhE) that first makes aldehyde and then ethanol. Ethanol may also be produced from pyruvate by pyruvate decarboxylase (PDC) coupled with a monofunctional ADH1. Pyruvate may also be converted to lactate by lactate dehydrogenase (LDH). CpLDH and CpMDH in C. parvum are better studied than many other glycolytic enzymes. These two genes are arranged in tandem in chromosome VII, and CpLDH likely evolved from CpMDH by a very recent gene duplication event after the separation of Cryptosporidium from other apicomplexans (Madern et al., 2004). In fact, based on strong phylogenetic evidence (Zhu and Keithly, 2002; Madern et al., 2004) all apicomplexan LDH and MDH genes were likely acquired from a α-proteobacterial ancestor. Unlike many other eukaryotes that possess several MDH isozymes (e.g., cytosolic, mitochondrial, or chloroplast), Cryptosporidium and even other mitochondrial- and plastid-containing apicomplexans appear to have only the cytosolic-type MDH, for which the significance in evolution and function has been discussed elsewhere (Madern et al., 2004). CpLDH, PK, and glyceraldehyde 3-phosphate dehydrogenase (GAPDH) have also been crystallized and their x-ray diffraction data collected for structural studies (Senkovich et al., 2005). The AceCL gene was reported 10 years ago (Khramtsov et al., 1996), but its biochemical features have not been studied so far. Cryptosporidium can also make N-glycans from fructose-6P or mannose (via mannose-6P), which is required for synthesizing some glycolipids. Enzymes involved in the synthesis of amylopectin and amylose from glucose and glucose-6P, including amylopectin phosphorylase, amylopectin 1,6-glucosidase, and the branching enzyme, are all identified in the C. parvum genome. Another unique feature is the ability to synthesize trehalose via glucose-1P and UDP-glucose, which is not seen in other apicomplexans. Trehalose is commonly found in a wide range of organisms, including bacteria, fungi, plants, and some invertebrates, and may function as an antidesiccant, antioxidant, or protein-stabilizing agent (Schmatz, 1989; Michalski et al., 1992). Because the mannitol cycle found in Eimeria spp. is not present in Cryptosporidium, it is assumed that trehalose may play a role similar to mannitol in Eimeria oocysts (Schmatz, 1989; Michalski et al., 1992). Whether trehalose can serve as an energy source for Cryptosporidium is questionable, because no trehalase ortholog responsible for breaking down trehalose can be identified from the parasite genomes. FIGURE 3.1 (A color version of this figure follows page 242.) Major metabolic pathways and their connections in Cryptosporidium based on genome annotation and recent biochemical data. Solid arrow bars indicate direct biochemical reactions, whereas dashed lines indicate multiple-step reactions or connections between pathways. Most enzymes are highlighted with solid dark green boxes, but those involved in ATP consumption or production are highlighted with orange and red boxes, respectively. Organic end products are boxed. Abbreviations: ABC, ATP-binding cassette transporter; ACBP, acyl-CoA binding protein; ACC, acetyl-CoA carboxylase; AceCL, acetate-CoA ligase (also termed acetyl-CoA synthetase); ACL, fatty acid-CoA ligase (also termed fatty acyl-CoA synthetase); ADC, arginine decarboxylase; ADH, alcohol dehydrogenase; adhE type E alcohol dehydrogenase; AIH, agmatine iminohydrolase; AK, adenosine kinase; AMPDA, AMP deaminase; dCMPDA, dCMP deaminase; CTPS, CTP synthase; DHF, dihydrofolate; DHFR, dihydrofolate reductase; CMPK, CMP kinase; CMPS, CMP synthase; GAPDH, glyceraldehyde phosphate dehydrogenase; GDH, glycerol phosphate dehydrogenase; GGH, glucoside glucohydrolase; GS, glutamine synthetase; HK, hexokinase; IMPDH, IMP dehydrogenase; LCE, long-chain fatty acyl elongase; LDH, lactate dehydrogenase; MDH, malate dehydrogenase; ME, malic enzyme; MPGT, mannose-1-phosphate guanylyltransferase; mTHF, methylene tetrahydrofolate; ORP, oxysterol-binding protein-related protein; PDC, pyruvate decarboxylase; PEPCL, phosphoenolpyruvate carboxylase; PFK, phosphofructokinase; PGI, phosphoglucose isomerase; PGK, phosphoglycerate kinase; PGM, phosphoglycerate mutase; PGluM, phosphoglucose mutase; PK, pyruvate kinase; PKS, polyketide synthase; PMI, mannose-6-phosphate isomerase; PMM, phosphomannomutase; PNO, pyruvate NADP+ oxidoreductase; PV, parasitophorous vacuole; PVM, parasitophorous vacuole membrane; RNR, ribonucleoside-diphosphate reductase; SHMT, serine hydroxymethyl transferase; SpdSyn, spermidine synthase; SpmSyn, spermine synthase; SSAT, spermidine:spermine N1-acetyltransferase; THF, tetrahydrofolate; TIM, triosephosphate isomerase; TIM17, translocase of inner mitochondrial membrane (17 kDa); TK, thymidine kinase; TOM40, translocase of outer mitochondrial membrane (40 kDa); TP, trehalose phosphatase; TPS, trehalose phosphate synthase; TS, thymidylate synthase; UDPGPP, UDP-glucose pyrophosphorylase; UDPK, UDP/CDP kinase; UK, uridine kinase; UMPK, UMP/CMP kinase; UPRT, uracil phosphoribosyltransferase.
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IV.
Cryptosporidium and Cryptosporidiosis, Second Edition
Nucleotide Metabolism
Similar to other apicomplexans, Cryptosporidium is unable to synthesize purines de novo. However, its purine scavenge pathway is highly streamlined and appears to solely rely on uptake of adenosine from host through a transporter (Figure 3.1). Adenosine is converted to AMP by an adenosine kinase (AK) (Galazka et al., 2006), which in turn is converted to IMP (by AMP deaminase, [AMPDA]), XMP (by IMP dehydrogenase [IMPDH]), and GMP (by GMP synthase [GMPS]). However, unlike other apicomplexans, Cryptosporidium lacks a gene encoding hypoxanthine-xanthine-guanine phosphoribosyltransferase (HXGPRT), indicating that GMP may only be made via a simple AMP-to-GMP pathway (Abrahamsen et al., 2004; Striepen and Kissinger, 2004). More surprisingly, CpIMPDH differs from other apicomplexan IMPDHs and is evolutionarily related to ε-proteobacterial homologues (Striepen et al., 2002; Umejiego et al., 2004). For this reason, CpIMPDH is much less sensitive to mycophenolic acid (MPA) than other eukaryotic homologues, including TgIMPDH. In fact, the CpIMPDH gene was first identified unexpectedly by complementation screening of a C. parvum expression library in a ΔHXGPRT strain of T. gondii (Striepen et al., 2002). The assay was originally designed to hunt for the hypothetical CpHXGPRT gene under the selection by MPA. More recently, this enzyme has been expressed as a recombinant protein, and its enzyme kinetics were characterized in detail (Umejiego et al., 2004). Additionally, CpIMPDH was also able to complement the function in an IMPDH-knockout strain of Escherichia coli, which not only confirms the function of this enzyme, but also makes it possible to perform bacterial growth-based high-throughput screening of anti-CpIMPDH compounds for drug development (Umejiego et al., 2004). Plasmodium and Toxoplasma (and probably many other apicomplexans) are capable of synthesizing pyrimidines de novo from glutamine (Gardner et al., 2002; Kim and Weiss, 2004). However, Cryptosporidium lacks a de novo pathway, and relies on nucleotide transporters for scavenging pyrimidines. This parasite makes UMP via two uracil phosphoribosyltransferases (UPRTs): one is a discrete enzyme, whereas the other is fused with uridine kinase (UK-UPRT). UK is also responsible for synthesizing CMP from cystidine, and thus is also termed as uridine-cystidine kinase. Cryptosporidium also possesses a bacterial-type thymidine kinase (TK) to synthesize dTMP that can be further converted to dTTP or dUMP, dCMP, and dCMP. The conversion from dTMP to dUMP is catalyzed by thymidylate synthase (TS), and coupled with folate metabolism. TS is fused with dihydrofolate reductase (i.e., bifunctional DHFR-TS) in all apicomplexans and many other protists. Cryptosporidium DHFR-TS is one of the few enzymes that have been more thoroughly studied and pursued as a drug target (Gooze et al., 1991; Vasquez et al., 1996; O’Neil et al., 2003; Lilien et al., 2004; Anderson, 2005a, 2005b). More recently, its crystal structure has been determined, which has revealed a unique linker domain that controls the relative orientation of the DHFR and TS domains (Anderson, 2005a).
V.
Fatty Acid Metabolism
Fatty acid biosynthesis consists of repeated elongations of 2-carbon (C2) units to a fatty acyl precursor (Figures 3.1 and 3.2). In de novo synthesis, the precursor is acetic acid or acetyl-CoA. Longer-chain fatty acids may also be used as precursors. Fatty acids are typically synthesized by fatty acid synthase (FAS), which actually represents a number of individual enzymes. Among them, acyl transferase (AT) first transfers the fatty acyl chain to an acyl carrier protein (ACP) to form fatty acyl-ACP (Smith, 1994). Newly translated ACP (i.e., apo-ACP) is not functional, and a prosthetic phosphopantetheine has to be added to a Ser residue to make functional holo-ACP (Figure 3.2) (Cai et al., 2005). The enzyme involved in activating ACP, referred to as phosphopantetheinyl transferase (PPT), has been identified from Cryptosporidium and some other apicomplexans. It turns out that there are two types of PPT correlated with the types of FAS in apicomplexans: Cryptosporidium has an SFP-type PPT responsible for activating Type I ACP; Plasmodium uses a plastid-specific ACPS-type PPT with plastid-targeting signal for Type II ACP, whereas Toxoplasma has both SFP- and ACPS-type PPTs (Table 3.1). CpSFP-PPT has been
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FIGURE 3.2 Domain organization and proposed reactions catalyzed by individual domains of CpFAS1 megasynthase. Targets for select inhibitors (triacsin C and cerulenin) are indicated by T-shaped bars. Abbreviations: ACP, acyl carrier protein; AL, acyl-[ACP] ligase; AT, acyl transferase; DH, dehydrase; ER, enoyl reductase; KR, ketoacyl reductase; KS, ketoacyl synthase; Ppant, phosphopantetheinyl moiety; PPT, phosphopantetheinyl transferase; Red, acyl-[ACP] reductase; R represents fatty acyl chain (saturated or unsaturated) with unspecified carbon lengths. However, the AL domain prefers medium- to long-chain fatty acids as its substrates. CpPKS1 (not shown here) contains seven internal modules and uses the same mechanism for polyketide chain elongation. Owing to the lack of one or more enzymatic domains within some modules, the keto groups may not be completely reduced and/or dehydrated, resulting in the presence of keto, hydroxyl, and/or enoyl bonds in the product.
TABLE 3.1 Correlation Between the Presence of Apicoplast and Types of Fatty Acid Synthase (FAS), Polyketide Synthase (PKS), and Phosphopantetheinyl Transferase (PPT) Among Different Groups of Apicomplexans Organism Cryptosporidium Plasmodium Toxoplasma Eimeria a
Apicoplast Absent Present Present Present
Type Type Type Type
Type of FAS or PKS
Type of PPTa
I FAS/PKS only II FAS only I FAS/PKS and Type II FAS I FAS/PKS and Type II FAS
SFP ACPS SFP and ACPS SFP and ACPS
ACPS = holo-acyl carrier protein synthase-type PPT; SFP = surfactin production element-type PPT.
cloned, and its biochemical features in activating the ACP domains in CpFAS1 and CpPKS1 in vitro have been thoroughly characterized (Cai et al., 2005). AT is also responsible for the formation of transient malonyl–ACP complex, before a C2 unit from a malonyl moiety is condensed into the fatty acyl-ACP via a decarboxylation process by ketoacyl synthase (KS). The keto group retained from the previous acyl chain is then reduced by ketoacyl reductase (KR), dehydrated by dehydrase (DH), and reduced again by enoyl reductase (ER) to yield a saturated fatty acyl chain (Figure 3.2). The elongation process may be repeated several times until the acyl chain reaches the desired length (e.g., C16 in typical de novo synthesis) (Smith, 1994). In humans, animals, and birds, these enzymes are fused into a single polypeptide that is cytosolic, multifunctional, and referred to as
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Type I FAS. In bacteria and plastid-containing eukaryotes, these enzymes are discrete and monofunctional, targeted to plastid in eukaryotes, and referred to as Type II FAS. Among apicomplexans, Plasmodium spp. possess only Type II FAS, whereas Toxoplasma and Eimeria have both Type I and II enzymes (Table 3.1). However, Cryptosporidium lacks an apicoplast and its associated Type II pathway, but possesses a 25-kb atypical Type I FAS (CpFAS1) consisting of at least 21 enzymatic domains (Figure 3.2) (Zhu et al., 2000d; Zhu et al., 2004). The loading unit contains acyl[ACP] ligase (AL) to load fatty acid to the adjacent ACP; three internal modules are assumed to perform at least three cycles of C2 elongation of the fatty acyl chain, and the C-terminal reductase domain may release the acyl chain as fatty aldehyde or alcohol using NAD(P)H as an electron donor (Figure 3.2). The release of acyl chain by reductase differs from that by thioesterase in typical FAS, which releases fatty acid via hydrolysis. Additionally, the genome of C. parvum also encodes a 45-kb polyketide synthase (CpPKS1) that resembles CpFAS1 but contains seven internal modules lacking one or more enzymatic domains (Zhu et al., 2002). Therefore, ketos, hydroxyl groups, and/or enoyl bonds may be retained in the products, a feature characteristic of the PKS family. The AL domains in CpFAS1 and CpPKS1 prefer long-chain fatty acids (LCFAs) or very LCFAs (VLCFAs) as substrates (Zhu et al., 2004; Fritzler and Zhu, 2007). Therefore, these two “megasynthases” are likely involved in the elongation of fatty acids or polyketides, rather than making fatty acids or polytides de novo, which differs from the Type I FASs in humans, animals, and fungi, or other typical microbial PKSs. The true products of CpFAS1 and CpPKS1, as well as their biological roles, remain to be determined. However, LCFAs and VLCFAs are present in C. parvum, detectable as simple or complex lipids in oocysts, or as an essential component in the highly antigenic glycoproteins in sporozoites (Mitschler et al., 1994; Priest et al., 2003). In addition to CpFAS1 and CpPKS1, Cryptosporidium also possesses a discrete long-chain fatty acyl elongase (CpLCE) that uses saturated or unsaturated fatty acyl-CoA as precursors (rather than acylACP) to make longer fatty acids (Zhu, 2004). Humans or yeasts each use three LCEs to make LCFAs (e.g., C16 and C18) or VLCFAs (e.g., C22 or C26) (Moon et al., 2001; Matsuzaka et al., 2002). More recently, it has been discovered that the LCE-associated pathway may be expanded and utilized to synthesize fatty acids de novo in trypanosomes (so not just a “long-chain” elongase in this case) (Lee et al., 2006). Similar to the FAS (acyl-ACP) system, the elongation of acyl-CoAs also comprises a serial reaction, in which LCE plays a role similar to KS; i.e., it is responsible for the first condensing reaction. There are also less well-characterized enzymes responsible for the subsequent keto-reduction, dehydration, and enoyl reduction. By possessing a single LCE, Cryptosporidium is unlikely to use it for synthesizing fatty acids de novo. Current molecular and biochemical data, including the lack of Type II FAS and the substrate preference toward LCFAs for CpFAS1, CpPKS1, and CpLCE, strongly indicate that Cryptosporidium is incapable of synthesizing fatty acids de novo (Zhu, 2004). How the parasite scavenges fatty acids from host cells and intestinal lumens is unclear. Both C. parvum and C. hominis genomes appear to lack homologues to any known eukaryotic or prokaryotic fatty acid transporters. Free fatty acids or simple lipids may simply diffuse across the membranes, or some of the uncharacterized membrane proteins may act as fatty acid/lipid transporters. On the other hand, the C. parvum genome possesses a P-type ATPase whose predicted substrate is phospholipid. Whether this protein is involved in transporting lipids is worth exploring. Additionally, some families of ATP-binding cassette (ABC) transporters are known to be involved in lipid transport, such as those in the peroxisomes (Theodoulou et al., 2006). Cryptosporidium possesses a large number of ABC transporters (see Section IV for details); thus, they may also utilize one or more of them to scavenge fatty acids or other lipids. Free fatty acids have to be activated to form acyl-ACP or acyl-CoA esters before they can enter subsequent metabolic pathways. In addition to FAS and PKS, which can directly activate fatty acids to form acyl-ACP by the AL domains, Cryptosporidium also has three discrete fatty acid-CoA ligases (ACLs) to make fatty acyl-CoAs. Because uncontained fatty acyl-CoA molecules may act as a detergent that is harmful to cell membranes, they have to be used immediately or stored or transported via an acyl-CoA binding protein (ACBP). Cryptosporidium has a single, “long-type” ACBP (CpACBP1) that is fused with an ankyrin-repeat sequence and prefers medium- to long-chain fatty acyl-CoAs as substrates (Zeng et al., 2006). CpACBP1 is chiefly localized to the parasitophorous vacuole membrane (PVM), rather than in the intracellular parasites or free sporozoites. Whether CpACBP1 is involved in the
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membrane remodeling in PVM or fatty acid transport across PVM remains to be elucidated. In addition to ACBP, Cryptosporidium also encodes two oxysterol-binding protein-related proteins (CpORP1 and 2), in which CpORP1 was also localized to PVM (Zeng and Zhu, 2006). Both CpORPs can bind to all species of phosphatidylinositol phosphates (PIPs). Whether and how CpORPs are involved in the regulation and transport of sterols or other lipids is not fully understood. However, the presence of ACB and ORP proteins in PVM indicates that this membrane barrier between parasite and intestinal lumen is directly involved in lipid transport and remodeling during the parasite’s intracellular development. Finally, Cryptosporidium lacks enzymes for β-oxidation of fatty acids into acetyl-CoA, which indicates that fatty acids are not an energy source for this parasite (if so, the product acetyl-CoA may be converted to acetate by AceCL to produce ATP). It also further reinforces the importance of glycolysis in the energy metabolism in Cryptosporidium.
VI. Polyamine Metabolism Polyamines are a group of small molecules essential to all cells. During polyamine synthesis, arginine is first converted to either ornithine by arginase, or agmatine by arginine decarboxylase (ADC). Ornithine or agmatine is further converted to putrescine by ornithine decarboxylase (ODC) or agmatine iminohydrolase (AIH). Longer polyamines, including spermidine and spermine, can be made from putrescine by spermidine or spermine synthase (SpdSyn or SpnSyn). However, spermine can be back-converted to spermidine, and to putrescine by a spermidine:spermine N-acetyltransferase (SSAT). Humans, animals, and fungi use the ODC pathway, whereas plants typically use ADC. Bacteria may have both ODC and ADC to make putrescine. Many protists use the ODC pathways. Among them, Trypanosoma ODC is the target for difluoromethylornithine (DFMO), an effective drug against African trypanosomiasis (Bacchi and Yarlett, 2002). Among apicomplexans, Cryptosporidium appears to possess only the plant-type ADC pathway, but not ODC (Figure 3.1) (Keithly et al., 1997). The ADC activity and C. parvum development could be inhibited by difluoromethylarginine (DFMA). However, P. falciparum differs from C. parvum (and many other apicomplexans) by possessing ODC that is fused with S-adenosylmethionine decarboxylase (SAMDC), rather than ADC (Krause et al., 2000; Muller et al., 2000; Wrenger et al., 2001; Birkholtz et al., 2004). The SAMDC ortholog is also absent in C. parvum. Preliminary analysis of the unpublished genome data indicates that E. tenella appears to encode a discrete ODC (as SAMDC and ODC orthologs are distributed at two different large contigs), whereas T. gondii may not be able to synthesize putrescine from arginine because it lacks both ODC and ADC (unpublished observations). However, although ADC activity was consistently detected in C. parvum, gene encoding ADC is not recognizable in its complete genome. It is possible that either the ADC gene is highly divergent in Cryptosporidium, or there is an alternative pathway that displays ADC-like activity in this parasite. The polyamine pathway has been pursued as a potential drug target in C. parvum. However, strong SSAT activities were also detected in this parasite (Figure 3.1), implying that an effective treatment can only be achieved when both forwardand back-converting pathways are targeted, along with the blockage of polyamine uptake from host cells or gut.
VII. Amino Acid Metabolism Although amino acids are the basic building blocks of proteins, Cryptosporidium apparently cannot synthesize any of them de novo. All amino acid synthetic genes are missing from the C. parvum genome. Instead, this parasite possesses at least 11 amino acid transporters for scavenging amino acids from host cells (and the intestinal lumen), in contrast to P. falciparum, which only possesses one amino acid transporter (Abrahamsen et al., 2004). However, C. parvum retains the capacity of interconverting a limited number of amino acids. Glutamate produced by GMP synthetase can be recycled back to glutamine (Gln) by glutamine synthetase (GS) (Figure 3.1). Serine (Ser) and glycine (Gly) may be
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interconverted by serine hydroxymethyl transferase (SHMT) within the folate metabolic pathway (Figure 3.1). Asparagine (Asn) can be made from aspartate (Asp) by asparagine synthetase, which might be important in the recycling of NH3 released by AMP deaminase. Other conversions include methionine (Met) and S-adenosylmethionine (SAM) by SAM synthetase, and homocysteine and S-adenosyl homocysteine (SAH) by SAH synthase. One small surprise is the presence of a single tryptophan synthase that may synthesize tryptophan (Trp) from indole or indoleglycerol phosphate within the Trp synthetic pathway.
VIII. Structural Proteins Structural proteins play critical roles in parasite cell biology and host–pathogen interactions. Similar to other apicomplexans, Cryptosporidium possesses a number of structures unique to the phylum Apicomplexa, including apical complex and oocyst wall. The unique shape of apicomplexan sporozoites and merozoites is maintained by the unique arrangement of cytoskeletal proteins (Morrissette and Sibley, 2002). Actins are one of the major components of the cytoskeleton. Most apicomplexans, including Cryptosporidium, possess only a single conventional actin (with the exception of Plasmodium, which has two) (Gordon and Sibley, 2005). However, Cryptosporidium also possesses seven actin-like proteins (Gordon and Sibley, 2005). Together with myosin, actin plays a critical role in sporozoite gliding motility in C. parvum and other apicomplexans. Recent studies have shown that actin- and myosin-dependent gliding motility is critical to the C. parvum sporozoite invasion. Inhibitors of actin and myosin could inhibit not only gliding motility, but also the invasion of C. parvum sporozoites into host cells (Chen et al., 2004; Sibley, 2004; Wetzel et al., 2005). Microtubules consisting of polymerized α- and β-tubulins are important in maintaining the parasite’s structure and cytokinesis. In addition to the common spindle microtubules that are typically formed during cell division, apicomplexans also have distinct subpellicular microtubules that radiate posteriad from the apical polar ring and cover about two-thirds of the body length near the surface of sporozoites or merozoites (Morrissette and Sibley, 2002). The Cryptosporidium β-tubulin gene contains only a single intron near the 5′ end (Caccio et al., 1997), which differs from its orthologs in other apicomplexans or eukaryotes that typically contain three introns (Zhu and Keithly, 1996). It is also interesting that the intron within the β-tubulin gene was not fully spliced out in its transcripts at later developmental stages in vitro (but normally spliced out in vivo) (Cai et al., 2004). It was speculated that the “deficiency” in removing the intron might be one of the factors that contributed to the inability of the parasite to complete the life cycle under in vitro conditions. The importance of β-tubulin (and actin) in the invasion of C. parvum into host cells has been recently demonstrated, as sporozoites treated with colchicines (and cytochalasin D) could inhibit apical organelle discharge and invasion (Chen et al., 2004). However, treatment by these two cytoskeletal inhibitors had no effect on the attachment of sporozoites to the host cell surface, indicating that cytoskeletal proteins may play little role in the parasite attachment. The oocyst wall plays a critical role in protecting Cryptosporidium sporozoites from environmental hazards. It is virtually resistant to all disinfectants at concentrations that may kill almost all other microbes. However, little is known regarding the composition and molecular structure of the oocyst wall. Cryptosporidium oocyst wall proteins (COWPs) from three Cryptosporidium species were first observed by SDS-PAGE analysis (Tilley et al., 1990). At least seven bands were shared among C. parvum, C. muris, and C. baileyi, and each species appeared to have one unique band. More reports were published later on, characterizing various COWPs based on specific antibodies (e.g., Bonnin et al., 1991; McDonald et al., 1991; Nina et al., 1992; Ranucci et al., 1993; Mead et al., 1994). The first COWP gene was initially reported as a 2.4-kb DNA fragment, and its entire open reading frame (ORF) was later determined to encode a 190-kDa protein that contains arrays of two types of cysteine-rich tandem repeats (designated as type I and type II domains) (Lally et al., 1992; Spano et al., 1997). Since then, the COWP gene has been extensively used as one of the genetic markers in genotyping various Cryptosporidium species and genotypes (Xiao et al., 2000; Akiyoshi et al., 2002; Kato et al., 2003). A 45-kDa COWP was later cloned and expressed (Jenkins et al., 1999). However, a more complete picture of COWPs was only obtained after the completion of the C. parvum genome sequencing project. The C. parvum genome in fact encodes
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at least nine COWPs (namely, from COWP1 to COWP9) (Templeton et al., 2004b). Type I domains are present in all COWPs, whereas Type II domains are only present in COWP1-3. All COWPs also possess an N-terminal signal peptide sequence for targeting them to the oocyst wall. As expected, all genes encoding these COWPs are expressed at elevated levels when oocyst formation is observed (i.e., at 48 to 72 h post infection). However, it is yet unclear how these COWPs are incorporated into the oocyst wall, as well as if there are other COWPs, because the search was largely based on the presence of the two types of domains. Besides COWPs, it is also unknown if (and what) other potential components (such as lipids and carbohydrates) may also contribute to the oocyst wall structure. A more recent study based on a lectin-binding assay has revealed the presence of N-acetyl-galactosamine-containing molecules on the surface of C. parvum oocysts, indicating that oocyst walls may contain more complex chemical structures than just proteins (Stein et al., 2006).
IX. Membrane Proteins and Transporters Cryptosporidium parvum surface proteins, including many species of glycoproteins and lipoproteins, have gained attention owing to their potential roles in parasite–host cell interactions and immune evasion. The molecular sizes of mucin-like glycoproteins on the surface of C. parvum (CpGPs) sporozoites can range from a few kDa up to 900 kDa (Tilley et al., 1991; Petersen et al., 1992; Petersen et al., 1997; Bonnin et al., 2001; O’Connor et al., 2002). In fact, over 20 genes are identified to encode mucin-like proteins, all of which contain hallmarks of extensive Thr or Ser stretches suggestive of O-glycosylation and signal peptide sequences conferring secretion (Templeton et al., 2004a). CpGPs are typical surface proteins, but some may be stored in micronemes before being translocated onto sporozoite surfaces (or even discharged in trails during gliding motility) (Tilley and Upton, 1994; Chen et al., 2004). One of the mucin-like proteins is GP40/15, which actually represents two proteins derived from a single 60kDa protein (also referred to as GP60) (Cevallos et al., 2000). Native GP40 could bind specifically to host cells, and antibodies against this protein could neutralize C. parvum infection in vitro, suggesting that this surface protein is involved in parasite adhesion and invasion (Cevallos et al., 2000). Both human furin and furin-like protease in C. parvum can cleave the 60-kDa protein at slightly varied sites (Wanyiri et al., 2006). The enzyme responsible for the activity is unknown. However, the C. parvum genome does have a subtilisin ortholog (EAK90066). Whether this subtilisin (or other protease) is involved in cleaving GP60 is an interesting and important subject to pursue because it may shed light on the general machinery of posttranslational cleavage of proteins. Another important finding related to GP60 is the successful expression of this protein in T. gondii, in which the heterogeneous protein is processed into GP40 and GP15 and glycosylated similarly as native antigens (i.e., by GalNAcα-O-Ser/Thr linker) (O’Connor et al., 2003). This observation, together with the successful complementation of CpIMPDH in T. gondii (Striepen et al., 2002; Striepen and Kissinger, 2004), indicates that the genetically tractable T. gondii system can serve as a surrogate for studying the function of proteins from Cryptosporidium for which no genetic system is yet available. Another group of smaller surface proteins (e.g., 3 to 27 kDa) are the glycosylphosphatidyl-inositol (GPI) anchored proteins, which might be important in immune evasion and pathogenesis (Priest et al., 2003). The dominant glycosylinositol phospholipid (GIPL) antigen contains a C18:0 lyso-acylglycerol, a C16:0-acylated inositol, and an unsubstituted mannose-3-glucosamine glycan core. Other diacyl species include a series of GIPLs containing an acyl-linked C20:0 to C28:0 lipid on the inositol ring. The presence of VLFAs suggests that C. parvum has to either scavenge these less common fatty acids from the host or synthesize them from common species such as C16 or C18 fatty acids, probably via CpFAS1or CpLCE-related pathways. Enzymes involved in GPI anchor synthesis can be readily identified from the C. parvum genome, such as phosphatidylinositol N-acetylglucosaminyltransferases (e.g., XP_628152, XP_627129 and XP_626317) (Templeton et al., 2004a). However, detailed biochemical features of GPI anchor synthesis in this parasite are as yet unexplored. As mentioned earlier, one unique feature of Cryptosporidium is the incapacity to synthesize almost anything de novo, which makes this parasite completely dependent on the host for its nutrients. Transporters for sugars, nucleotides, and amino acids can be readily identified from the genome (Abrahamsen
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et al., 2004; Xu et al., 2004), although their detailed substrate specificity and kinetics remain to be determined. Additionally, Cryptosporidium also possesses about 19 ABC transporters that are involved in transporting various molecules across membranes (e.g., metabolites, lipids and sterols, and drugs) (Abrahamsen et al., 2004), one of which has been previously localized at the host cell–parasite boundary, reinforcing the importance of electron-dense structure (the feeder organelle) in the transport of substances between host cell and parasite (Perkins et al., 1999). A few more were also cloned, and their distribution in sporozoites was determined (Zapata et al., 2002). At least four genes that encode ABC proteins belong to the sbmA family. Members of this family are typically involved in the transport of peptide antibiotics in bacteria or multidrug resistance in a number of pathogens or cancer cells. Whether these pumps are involved in the resistance of this parasite to some drugs is unclear. Another gene of interest encodes a PfCRT ortholog responsible for chloroquine resistance in malarial parasites. The function of these transporters remains to be elucidated. The C. parvum genome also encodes at least seven P-type ATPases (CpATPases) that are typically involved in transporting cations. However, at least one P-ATPase from C. parvum belongs to the phospholipid group, which may be involved in lipid remodeling or transport. Three CpATPases have been reported, including a putative Ca2+-ATPase that is chiefly localized near the nuclear and apical regions (Zhu and Keithly, 1997); a heavy metal transporter that specifically binds reduced copper, Cu(I) (LaGier et al., 2001); and one that belongs to a group whose substrates are yet unknown (LaGier et al., 2002). Another ion pump is the vacuolar proton translocating ATPase (V-type ATPase). Genes encoding all subunits of the V-type ATPase are identified in the parasite genome, which includes the largest subunit with seven transmembrane regions (Abrahamsen et al., 2004). The subcellular location of this proton pump is yet to be determined, so it is not known whether this proton pump functions in lysosomes or other locations in this parasite. Cryptosporidium also possesses a total of 12 genes that encode thrombospondin-related adhesive proteins (TRAPs) that contain one or more thrombospondin Type 1 (TSP1) domains (Deng et al., 2002). TRAPs have been found in all apicomplexans examined so far. A series of studies have revealed that TRAPs may play a critical role in mediating host–parasite interactions and parasite gliding motility (Sultan et al., 1997; Wengelnik et al., 1999; Kappe et al., 2003; Kappe et al., 2004; Baum et al., 2006). Disruption of the TRAP gene in P. falciparum can block parasite motility and reduce infectivity, suggesting that TRAP might serve as a candidate gene for vaccine development (Dolo et al., 1999). Because there are multiple TRAPs in C. parvum, it is questionable whether these proteins can be effectively pursued as targets for immunological intervention against cryptosporidiosis. An early study using monoclonal antibodies to C. parvum TRAPC1 functional domains demonstrated limited efficacy against development in vitro (Camero et al., 1999). The presence of 12 TRAP genes indicates that Cryptosporidium possesses a complex and redundant machinery to sustain gliding motility and interactions with host cells.
X.
DNA and RNA Metabolism
Because of the lack of extrachromosomal DNA, Cryptosporidium only needs to maintain a single nuclear genome. Few studies have delineated DNA replication, repair and recombination, or gene activation and transcription, which are important in dissecting the machinery governing the unique life and cell cycles of this parasite. However, Cryptosporidium does utilize typical eukaryotic machinery (e.g., including typical eukaryotic DNA and RNA polymerases) for DNA replication and transcription (Abrahamsen et al., 2004). The components involved in these processes are considerably reduced. Genes encoding α-, β-, and ε-subunits of DNA polymerases are present, along with other key elements such as ORC1-5, MCMs, RF-A, RF-Cs, and CDC45. Oocysts may be exposed to extreme environmental factors, including UV irradiation, that cause DNA damage. Cryptosporidium apparently possesses a set of DNA repair proteins (including UV-induced factors) to repair such damage, which include the η-subunit of DNA polymerase (involved in error-prone repair), a complete excision repair apparatus (Ercc4/XPF and XPG and the accessory components such as XPA and Ercc1). However, as the ability to resist UV damage
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in every organism is limited, UV irradiation at appropriate levels is still an effective way to inactivate oocysts (Rochelle et al., 2004; Rochelle et al., 2005; Al-Adhami et al., 2007). Because Cryptosporidium has a sexual stage in its life cycle, whether and how often recombination occurs in this parasite is a subject important for parasite biology and speciation based on the concept of reproductive separation of biological species. The availability of a large number of genetic markers for genotyping strains or species has a great potential to facilitate study in this area. Interspecific recombination under both experimental and natural conditions has been reported (Widmer, 2004). This is congruent with the presence of an archaeal-type topoisomerase VI (which might function similar to eukaryotic Spo11 protein in meiosis), and at least two RecA homologues, including DMC1, involved in meiotic recombination in the Cryptosporidium genome (Abrahamsen et al., 2004). Cryptosporidium possesses two distinct types of replication protein A large subunits (RPA1), one of which belongs to the short-type proteins only found in unicellular organisms (Zhu et al., 1999; Millership and Zhu, 2002). All three RPA subunits (i.e., RPA1 to 3) have been identified from the genome, and some of their biochemical features, including DNA-binding kinetics and dissociation constants, have been studied in detail (Millership et al., 2004a; Rider et al., 2005). RPA is actually a single-stranded DNA (ssDNA)-binding protein (SBP) involved in DNA replication, repair, and recombination. The presence of unique RPA species in Cryptosporidium [and other apicomplexans or some distant protists such as the trypanosomatid Crithidia fasciculate (Brown et al., 1994; Voss et al., 2002)] implies a possible significant difference between apicomplexans and higher eukaryotes in regulating DNA replication, repair, or recombination. Cryptosporidium only has one TATA-binding protein (TBP) as part of the basal transcription machinery (Millership et al., 2004b). However, the TBP binding sequences within this AT-rich genome are not known. Transcription activators may directly recruit TBP to the promoter region of the gene to be activated. In some cases, a transcriptional co-activator may be involved, such as multiprotein bridging factor 1 (MBF1), which is involved in cell differentiation or stress responses (Takemaru et al., 1997; Takemaru et al., 1998; Kabe et al., 1999). The C. parvum ortholog of MBF1 was cloned, and its function was confirmed by yeast complementation assay and by its ability to interact with CpTBP (Zhu et al., 2000b; Millership et al., 2004b). However, genes regulated by CpMBF1 have not been identified, and further studies are needed to dissect its biological roles in parasite development. RNA splicing factors are present in C. parvum, but in significantly reduced numbers in comparison with those of Plasmodium, which may correlate with the much-reduced intron contents in the genome (Abrahamsen et al., 2004). Intron containing mRNA in C. parvum cultured in vitro has been first observed in the β-tubulin transcripts (Cai et al., 2004) and, later, among other gene transcripts (unpublished observation). The biological significance remains to be determined. A negative observation worth mentioning is the lack of key components involved in small RNA-mediated posttranscriptional gene silencing (e.g., RNA-dependent RNA polymerase, Argonaute and DICER orthologs) (Abrahamsen et al., 2004). Consequently, RNAi-related technologies cannot be used to study gene function in this parasite. However, Cryptosporidium does possess partitivirus-like double-stranded RNAs (dsRNA) that encode an RNAdependent RNA polymerase and a protein that probably functions as the capsid (Khramtsov et al., 1997; Khramtsov and Upton, 2000). The viruslike dsRNA appears to be present in almost all Cryptosporidium species, and has been used as a genetic marker for detection and genotyping analysis (Xiao et al., 2001; Leoni et al., 2003, 2006). Individual viral particles have never been observed. However, if they can be isolated and manipulated, they might be used as a transfection vector in Cryptosporidium.
XI. Potential Drug Targets Currently, no completely effective drugs are available to treat cryptosporidiosis in humans and animals. Therefore, one of the major goals of ongoing and future studies on Cryptosporidium biochemistry is to identify and validate potential molecular targets for the development of new drugs against cryptosporidiosis in humans and animals, particularly for treating infections in immunocompromised patients. The complete C. parvum genome has revealed the extreme evolutionary and biochemical divergence of Cryptosporidium from other apicomplexans, including intestinal coccidia. Indeed, many proven or
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promising drug targets found in other apicomplexans are either absent (e.g., HXGPRT, ODC, mannitol cycle, shikimate pathway, electron transport chain, and apicoplast) or highly divergent (e.g., IMPDH), which explains why this parasite is insensitive to the compounds that are usually effective against other apicomplexans. However, the genome does encode many potential drug targets. Unique targets include (but not limited to): 1. Plant- or bacterial-type enzymes, including PPi-PFK, LDH, MDH, PEPCL, PGI, PNO, IMPDH, adhE, and TK. A more complete list of genes with apparent bacterial origins was reported elsewhere (Huang et al., 2004). 2. Proteins that are highly divergent or absent in humans and animals, including the giant modular CpFAS1 and CpPKS1, plus AOX, DHFR-TS, and enzymes involved in the synthesis of treholose and amylopectin. 3. Cytokeletal proteins that display unique kinetics in polymerization, including actins and βtubulin. Polyamine enzymes are another group of drug targets. However, studies on this pathway have been greatly hindered by the inability to identify the ADC gene in the parasite genome, as well as the strong back-converting activity. Classic drug targets may also be pursued for new drug development by studying the action of wellcharacterized drugs, such as nitazoxanide (NTZ) and paromomycin. Cryptosporidium lacks a typical PFO to serve as a target for NTZ. Therefore, the unique PFO domain-containing PNO may act on NTZ by reducing its nitro group to form a biotoxic radical, but the notion still remains to be validated experimentally (Coombs and Muller, 2002). Paromomycin acts on the aminoacyl tRNA site of ribosomes or the maturation of tRNA that has been overlooked in C. parvum (Lynch and Puglisi, 2001; Tekos et al., 2003; Abelian et al., 2004; Kaul et al., 2006). Although paromomycin is not highly effective against cryptosporidial infection in vivo, studying the mechanism of action of paromomycin on the aminoacyl tRNA may facilitate the development of new compounds with better efficacies.
XII. Some Technical Perspectives Related to Biochemical Studies Biochemical analysis of C. parvum proteins relies largely on the successful expression and purification of recombinant proteins. Significant progress has been made to overcome some technical difficulties in expressing the AT-rich genes in bacteria, which includes the use of bacterial strains containing extra sets of tRNA genes to correct the codon usage bias between eukaryotic and bacterial genes (e.g., Rosetta from Novagen or BL21-CodonPlus from Stratagene) and induction of expression at lower temperature (as low as 16˚C) to increase the solubility of fusion proteins (e.g., Millership and Zhu, 2002; Millership et al., 2004a; Zhu et al., 2004; Rider et al., 2005). In some cases, satisfactory expression could be achieved only by using artificially synthesized genes based on bacterial codon preference (Cai et al., 2005; Galazka et al., 2006). Maltose-binding protein (MBP)-based pMAL vectors (New England Biolabs) are the most successful system for expressing various Cryptosporidium proteins, particularly large proteins. However, other systems such as T7 RNA polymerase-based pET vectors with various tags (Novagen) and glutathione-S-transferase (GST)-based fusion systems are also commonly used for relatively small proteins (Zhu and Keithly, 1997; LaGier et al., 2001; LaGier et al., 2002; Cai et al., 2005). Large quantities of protein can be obtained using the baculovirus expression system in insect cells (Iochmann et al., 1999), which is a little more complex than bacterial expression systems, but may produce better folded and posttranslationally modified fusion proteins necessary for the function of many eukaryotic proteins. Other eukaryotic systems that have been explored to express C. parvum proteins include Saccharomyces cerevisiae and T. gondii (Brophy et al., 2000; Zhu et al., 2000b; Striepen et al., 2002; LaGier et al., 2003; O’Connor et al., 2003; Slapeta and Keithly, 2004). However, relatively small amounts of recombinant proteins have been isolated from these systems. Therefore, these systems have been used primarily to study the targeting and cell biology, or functional complementation of C. parvum
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genes or proteins. There are a few other systems available today that can produce a large amount of fusion proteins (e.g., yeast Pichia pastoris or microalgae Chamydomonas reinhardtii expression systems), which may be explored for expressing bioactive parasite proteins. Biochemical study and drug discovery are still hampered by difficulties. The inability to obtain and purify sufficient quantities of intracellular stages in vitro, particularly sexual stages and developing oocysts, makes it extremely difficult (if not impossible) for biochemical analysis of parasite development. A genetic system is not available for Cryptosporidium, which makes it impossible to truly validate the drug targets (by gene knockout or replacement) or ultimately dissect the gene functions and protein cell biology. Nonetheless, with the availability of whole-genome sequences, the next breakthroughs in C. parvum research will likely be the development of an effective genetic system and the emergence of novel drugs to treat human and animal cryptosporidiosis.
Acknowledgments Studies derived from the author’s laboratory have been mainly supported by grants from the National Institute of Allergy and Infectious Diseases (NIAID) at the National Institutes of Health (NIH) (R01 AI44594 and R21 AI055278).
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Wetzel, D.M., Schmidt, J., Kuhlenschmidt, M.S., Dubey, J.P., and Sibley, L.D. 2005. Gliding motility leads to active cellular invasion by Cryptosporidium parvum sporozoites. Infect. Immun. 73, 5379–5387. Widmer, G. 2004. Population genetics of Cryptosporidium parvum. Trends Parasitol. 20, 3–6. Wrenger, C., Luersen, K., Krause, T., Muller, S., and Walter, R.D. 2001. The Plasmodium falciparum bifunctional ornithine decarboxylase, S-adenosyl-L-methionine decarboxylase, enables a well balanced polyamine synthesis without domain-domain interaction. J. Biol. Chem. 276, 29651–29656. Xiao, L., Limor, J., Morgan, U.M., Sulaiman, I.M., Thompson, R.C., and Lal, A.A. 2000. Sequence differences in the diagnostic target region of the oocyst wall protein gene of Cryptosporidium parasites. Appl. Environ. Microbiol. 66, 5499–5502. Xiao, L., Limor, J., Bern, C., and Lal, A.A. 2001. Tracking Cryptosporidium parvum by sequence analysis of small double-stranded RNA. Emerg. Infect. Dis. 7, 141–145. Xu, P., Widmer, G., Wang, Y., Ozaki, L.S., Alves, J.M., Serrano, M.G., Puiu, D., Manque, P., Akiyoshi, D., Mackey, A.J., Pearson, W.R., Dear, P.H., Bankier, A.T., Peterson, D.L., Abrahamsen, M.S., Kapur, V., Tzipori, S., and Buck, G.A. 2004. The genome of Cryptosporidium hominis. Nature 431, 1107–1112. Zapata, F., Perkins, M.E., Riojas, Y.A., Wu, T.W., and Le Blancq, S.M. 2002. The Cryptosporidium parvum ABC protein family. Mol. Biochem. Parasitol. 120, 157–161. Zeng, B., Cai, X., and Zhu, G. 2006. Functional characterization of a fatty acyl-CoA-binding protein (ACBP) from the apicomplexan Cryptosporidium parvum. Microbiology 152, 2355–2363. Zeng, B. and Zhu, G. 2006. Two distinct oxysterol binding protein-related proteins in the parasitic protist Cryptosporidium parvum (Apicomplexa). Biochem. Biophys. Res. Commun. 346, 591–599. Zhu, G. and Keithly, J.S. 1996. The beta tubulin gene of Eimeria tenella. Mol. Biochem. Parasitol. 76, 315–319. Zhu, G. and Keithly, J.S. 1997. Molecular analysis of a P-type ATPase from Cryptosporidium parvum. Mol. Biochem. Parasitol. 90, 307–316. Zhu, G., Marchewka, M.J., and Keithly, J.S. 1999. Cryptosporidium parvum possesses a short-type replication protein A large subunit that differs from its host. FEMS Microbiol. Lett. 176, 367–372. Zhu, G., Keithly, J.S., and Philippe, H. 2000a. What is the phylogenetic position of Cryptosporidium? Int. J. Syst. Evol. Microbiol. 50, 1673–1681. Zhu, G., LaGier, M.J., Hirose, S., and Keithly, J.S. 2000b. Cryptosporidium parvum: Functional complementation of a parasite transcriptional coactivator CpMBF1 in yeast. Exp. Parasitol. 96, 195–201. Zhu, G., Marchewka, M.J., and Keithly, J.S. 2000c. Cryptosporidium parvum appears to lack a plastid genome. Microbiology 146, 315–321. Zhu, G., Marchewka, M.J., Woods, K.M., Upton, S.J., and Keithly, J.S. 2000d. Molecular analysis of a Type I fatty acid synthase in Cryptosporidium parvum. Mol. Biochem. Parasitol. 105, 253–260. Zhu, G. and Keithly, J.S. 2002. Alpha-proteobacterial relationship of apicomplexan lactate and malate dehydrogenases. J. Eukaryot. Microbiol. 49, 255–261. Zhu, G., LaGier, M.J., Stejskal, F., Millership, J.J., Cai, X., and Keithly, J.S. 2002. Cryptosporidium parvum, the first protist known to encode a putative polyketide synthase. Gene 298, 79–89. Zhu, G. 2004. Current progress in the fatty acid metabolism in Cryptosporidium parvum. J. Eukaryot. Microbiol. 51, 381–388. Zhu, G., Li, Y., Cai, X., Millership, J.J., Marchewka, M.J., and Keithly, J.S. 2004. Expression and functional characterization of a giant Type I fatty acid synthase (CpFAS1) gene from Cryptosporidium parvum. Mol. Biochem. Parasitol. 134, 127–135.
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4 Epidemiology
Gordon Nichols
CONTENTS I. Introduction ............................................................................................................................. 80 II. Surveillance ............................................................................................................................. 80 III. Incubation Period .................................................................................................................... 80 IV. Asymptomatic Infection.......................................................................................................... 81 V. Duration of Symptoms............................................................................................................ 81 VI. Molecular Epidemiology......................................................................................................... 82 VII. Seroepidemiology/Population Immunity ................................................................................ 85 VIII. Infants and Children................................................................................................................ 86 IX. Immunocompromised Populations.......................................................................................... 86 X. Age and Sex ............................................................................................................................ 86 XI. Age and Susceptibility ............................................................................................................ 86 XII. Differences in the Epidemiology of Cryptosporidium spp. ................................................... 87 XIII. Risk Factors............................................................................................................................. 87 XIV. Sources .................................................................................................................................... 87 XV. Transmission Routes ............................................................................................................... 89 XVI. Drinking Water ........................................................................................................................ 90 XVII. Other Drinking Waters ............................................................................................................ 92 XVIII. Recreational Water .................................................................................................................. 92 XIX. Person-to-Person Transmission ............................................................................................... 93 XX. Zoonotic Transmission ............................................................................................................ 93 XXI. Food and Drink ....................................................................................................................... 93 XXII. Travel....................................................................................................................................... 94 XXIII. Protection of Vulnerable Populations ..................................................................................... 95 XXIV. Exclusions on the Basis of Infection...................................................................................... 95 XXV. Geographic Distribution.......................................................................................................... 95 XXVI. Seasonality............................................................................................................................... 95 XXVII. Burden of Disease ................................................................................................................... 96 XXVIII. Descriptive and Analytical Investigations............................................................................... 97 XXIX. Sporadic Disease ..................................................................................................................... 97 XXX. Period Prevalence .................................................................................................................... 98 XXXI. Mixed Infections ..................................................................................................................... 98 XXXII. Human Volunteer Studies........................................................................................................ 98 XXXIII. Oocyst Viability, Infectivity, Quality, and ID50 ..................................................................... 98 XXXIV. Risk Assessment...................................................................................................................... 99 XXXV. Intervention.............................................................................................................................. 99 XXXVI. Conclusions ............................................................................................................................. 99 References........................................................................................................................... 101
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I.
Cryptosporidium and Cryptosporidiosis, Second Edition
Introduction
Cryptosporidiosis in humans is predominantly a diarrheal disease, occurring in all age groups, with the organisms growing in an intracellular but extracytoplasmic location in the enterocytes of the gastrointestinal tract. The disease is more protracted, severe, and affects extraintestinal sites in people with innate or acquired deficiencies in immunity (Hunter and Nichols, 2002). The clinical disease and pathology are presented in Chapter 8. Cryptosporidium causing symptomatic disease was first noted in turkeys in 1955. During the 1970s Cryptosporidium infections were reported to cause neonatal diarrhea in calves (Barker and Carbonell, 1974; Meuten et al., 1974; Morin et al., 1976; Pohlenz et al., 1978). The first human cases of cryptosporidiosis in humans were recorded in the 1970s: one in a young girl with enterocolitis (Nime et al., 1976) and the other in an AIDS patient (Meisel et al., 1976). With the developing AIDS pandemic in the 1980s more cases of cryptosporidiosis in AIDS were identified (Ma and Soave, 1983; Navin and Juranek, 1984; Pitlik et al., 1983; Soave et al., 1984), and Cryptosporidium was found to be a cause of diarrhea in immunocompetent people (Baxby et al., 1983; Blagburn and Current, 1983; Current et al., 1983; Horen, 1983; Pitlik et al., 1983; Reese et al., 1982; Tzipori, 1983). Cryptosporidiosis occurs worldwide. Seroepidemiological studies of particular areas have indicated that the percentage of the population affected at some time in life can vary from under 20% to over 90% (Dillingham et al., 2002).
II.
Surveillance
Surveillance provides the foundation for understanding the epidemiology of cryptosporidiosis (Nichols, 2003). Good laboratory detection coupled with timely reporting and regional or national analysis and reporting are essential for detecting all but the largest of outbreaks. Without routine laboratory detection of cases of cryptosporidiosis from among patients with diarrhea, it is unlikely that most outbreaks will be detected. Surveillance provides basic descriptive information about the disease, usually including the age, sex, date of illness, geographic location, and other demographic features, together with any information about the source and route of transmission of the infection. Surveillance information differs between and within countries and can be affected by reporting practices and social changes as well as patterns of disease. Some of the features of surveillance of waterborne diseases and cryptosporidiosis have recently been reviewed (Craun et al., 2006a; Nichols et al., 2006). The quality of surveillance data can be influenced by managed-care medical systems; by the reduced access to medical treatment at weekends, and on national holidays; by selection criteria applied to testing fecal samples in the laboratory; by the sensitivity and specificity of the diagnostic tests; by the timeliness and completeness of reporting to surveillance; and by the percentage of samples sent for reference (Nichols et al., 2006).
III. Incubation Period Cryptosporidiosis has an incubation period of around 5 to 7 days. The incubation period is based on studies in which the time of exposure is known and is based on the time or date of onset of symptoms and signs of infection (in animals the prepatent period is based only on signs). During the large drinking-water-related outbreak in Milwaukee, the mean incubation period was estimated to range from 3 to 7 days. A study of emergency room admissions of elderly people during the Milwaukee outbreak suggests an incubation period of 5 to 6 days based on measurements of when the water was turbid (Naumova et al., 2003), which is lower than that seen for children (7 days) and adults (8 days) (Morris et al., 1998). An estimation of incubation periods for a swimming pool outbreak suggested people were ill 5 days after exposure (Insulander et al., 2005). In a large waterborne outbreak in Japan, there were 14 people who had a time-limited exposure to the contaminated drinking water, and the mean incubation period was 6.4 days (range 5 to 8). A restaurant outbreak in Spokane, WA,
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resulted in incubation periods of between 3 and 9 days (Anon., 1998). In an outbreak linked to apple cider, the incubation period was 6 days (range: 10 hours to 13 days). An outbreak of cryptosporidiosis in HIV-positive patients through contaminated ice found a longer incubation period (13 days) compared to that previously reported for people without an immunodeficiency (Ravn et al., 1991). There is clearly quite a degree of variation in the time from exposure to symptoms among patients in an outbreak. Although some of this variation probably results from inaccuracies in the estimation of exposure events and symptom onset, there also appear to be genuine differences in the time required to get symptoms. There is a possibility that prior infection might alter the incubation period, and the inoculum size might similarly influence the incubation period, although evidence for this is sparse. The extent to which the incubation period varies between different species of Cryptosporidium remains unclear.
IV.
Asymptomatic Infection
A study compared symptomatic and asymptomatic cryptosporidiosis in 13 studies in the Nordic countries (Denmark, Finland, Norway, and Sweden) (Horman et al., 2004). The Cryptosporidium prevalence in asymptomatic people was 0.99% (0.81; 1.19) compared to 2.91% (2.71; 3.12) in patients with symptoms. In developing countries with poor sanitation, asymptomatic carriage can be high, with a survey in Bolivia finding that 31.6% of people carried the organism (Esteban et al., 1998). Asymptomatic infection is therefore probably related to exposure to viable oocysts. The extent of asymptomatic infection may be underdetected because current microscopic methods have limited sensitivity compared to molecular ones for direct detection of the organism in feces (Balatbat et al., 1996). Endoscopy has been used in assessing asymptomatic carriage (Doganci et al., 2002). Studies of volunteers infected with Cryptosporidium show that infected people develop antibody responses to 27-, 17-, and 15-kDa C. parvum antigens. Increases in specific antibody reactivity measured by immunoblot were more prevalent among volunteers who developed symptoms of cryptosporidiosis than among asymptomatic infected or oocyst-negative volunteers (Moss et al., 1998b). Volunteers with preexisting IgG antibody to the 27-kDa antigen excreted significantly fewer oocysts than volunteers without this antibody. IgG antibodies to the 17-kDa antigens and IgM to the 27-kDa antigens were higher at the time of exposure for asymptomatic infected persons than for those who developed symptoms. These results suggest that characteristic antibody responses develop following C. parvum infection and that persons with preexisting antibodies may be less likely to develop illness (Moss et al., 1998b). It remains difficult to determine the degree to which immunity plays an important role in asymptomatic disease in a community. If asymptomatic infection is occurring commonly, then oocysts from this population may be a more important source of oocysts in sewage than symptomatic patients.
V.
Duration of Symptoms
The duration of symptoms for people infected with Cryptosporidium is relatively easily determined for people identified with symptomatic cryptosporidiosis. However, people who do not consult a general practitioner may be less severely ill and be symptomatic for a shorter period. When a questionnaire is being completed during outbreak investigations, the symptoms may not be over at the time the questions about onset and duration are asked, and this might underestimate the duration of symptoms. During outbreaks the symptomatic period has ranged from 5 to 9 days (range 2 to 13) (Insulander et al., 2005; Lim et al., 2004). The duration of symptoms has been measured in hours in human volunteer studies. The duration in four different isolates of C. parvum ranged from 6 to 360 h, with differences between isolates not being significant (Okhuysen et al., 1999, 2002). Long-term sequelae resulting from infection can be common (Hunter et al., 2004a) and can cause nonintestinal symptoms. It has been reported that these symptoms are more commonly associated with
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C. hominis than with C. parvum (Hunter et al., 2004a). In people with a profound immunodeficiency, there can be symptoms that are different and more prolonged, and disease can result in an earlier death than would be the case if the patient had not been infected.
VI. Molecular Epidemiology The development of molecular methods for the sensitive detection, genotyping, and subgenotyping over the last few years has allowed the identification of sources of oocyst contamination and routes of transmission in both outbreak and nonoutbreak situations to be examined (Xiao et al., 2004a; Xiao and Ryan 2004; see Chapter 5 for details). The methods have developed from Western blotting approaches undertaken in England and Wales in the late 1980s (McLauchlin et al., 1998; Nichols et al., 1991, 1992) that demonstrated phenotypic differences between isolates. They have subsequently been superseded by PCR and sequencing approaches that have been used for both species differentiation and subtyping (McLauchlin et al., 1998, 1999, 2000; Nichols and McLauchlin, 2003; Patel et al., 1998, 1999; Peng et al., 1997; Spano et al., 1997, 1998a, 1998b). The PCR methods initially separated isolates into Genotype 1 (since renamed C. hominis) (Morgan-Ryan et al., 2002) and Genotype 2 (now called C. parvum). The approach has allowed molecular methods to be improved in sensitivity and allowed greater discrimination between isolates. Methods for the discrimination of isolates between and within species have used a variety of genotyping and subtyping tools. Genetic analysis has included PCR-RFLP and sequence analysis of the SSU rRNA (Xiao et al., 2004b), heat shock protein (HSP70) (Gobet and Toze, 2001; Sulaiman et al., 2000), the 60kDa glycoprotein (GP60) gene (Abe et al., 2006; Peng et al., 2003; Cohen et al., 2006; Leav et al., 2002; Muthusamy et al., 2006), Cryptosporidium oocyst wall protein (COWP) (McLauchlin et al., 1999; Patel et al., 1998), and small double-stranded viruslike RNA (Leoni et al., 2006c; Xiao et al., 2001b). Microsatellite and minisatellite analysis (Aiello et al., 1999; Caccio et al., 2000; Mallon et al., 2003a; Tanriverdi and Widmer, 2006) and multilocus sequence typing (Gatei et al., 2007) have also been used recently. The sensitivity of different PCR methods varies. Molecular methods have the advantage of an increased sensitivity of detection, particularly in immunocompromised patients (McLauchlin et al., 2003; Rodrigues et al., 2004). The methods have allowed a wider variety of species of Cryptosporidium to be identified in human feces, and have indicated limitations regarding the possible sources of oocysts. Typing has proved useful in case-control studies of sporadic Cryptosporidium infection (Goh et al., 2004; Hunter et al., 2004b), and also has proved useful in examining outbreaks (McLauchlin et al., 1998; Nichols and McLauchlin, 2003; Patel et al., 1998; Peng et al., 1997). Subtyping has shown identity between isolates from water and patients (Zhou et al., 2003). It is likely that the number of identified species causing disease in animals will increase and that, along with this, the number causing human disease will likely rise. Some of the species named in the past have not been validated with molecular methods. Many of the species contain a number of distinct genotypes and there are animal-specific genotypes that have yet to be named as distinct species. In an examination of over 13,000 fecal samples in England and Wales between 1989 and 2005, the percentage of human cases caused by C. parvum and C. hominis is approximately equal (Table 4.1) (Nichols et al., 2006). C. meleagridis made up less than 1% of cases, and genotypes other than C. parvum or C. hominis made up only 3.6%. C. parvum has a number of subtypes that can be grouped into subtype families as measured by GP60 sequence analysis (Alves et al., 2006). The occasional occurrence of C. hominis in cattle and goats (Park et al., 2006) means that careful comparisons of human and animal strains are required to elucidate the epidemiology of human cryptosporidiosis. There is evidence of human infection with C. felis (Caccio et al., 2002; Caccio 2005; Cama et al., 2003; Chacin-Bonilla, 2001, 2002; Coupe et al., 2005; Gatei et al., 2002b; Guyot et al., 2001, 2002; Joachim, 2004; Leoni et al., 2003, 2006a; Lindergard et al., 2003; Matos et al., 2004; Muthusamy et al., 2006; Pedraza-Diaz et al., 2001a; Pieniazek et al., 1999; Tiangtip and Jongwutiwes, 2002; Xiao et al., 2001a), C. meleagridis (Akiyoshi et al., 2003; Caccio 2005; Coupe et al., 2005; Enemark et al., 2002; Gatei et al., 2002b, 2003; Glaberman et al., 2001; Guyot et al., 2001, 2002; Leoni et al., 2003,
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83 TABLE 4.1 Cryptosporidium Species/Genotypes Detected Among 13,112 Human Cases in England and Wales 1989–2005 Cryptosporidium Species C. hominis C. parvum C. hominis and C. parvum C. meleagridis C. felis C. canis C. suis Cervine genotype Skunk genotype CZB141 genotype Novel or undetermined species/genotype (under further investigation)
Number of Patients 6594 5981 65 99 22 2 1 6 1 1 337
(50.29%) (45.6%) (0.5%) (0.8%) (0.2%) (0.02%) (0.01%) (0.05%) (0.01%) (0.01%) (2.6%)
Source: From Anon. 2002. The development of a national collection for oocysts of Cryptosporidium. Final Report to DEFRA, Drinking Water Inspectorate. Foundation for Water Research, Marlow, Bucks, U.K. (http://www.fwr.org/); Chalmers, R.M. et al., 2002a. Infection with unusual types of Cryptosporidium is not restricted to immunocompromised patients. J. Infect. Dis. 185, 270–271; Leoni, F. et al., 2006b. Genetic analysis of Cryptosporidium from 2414 humans with diarrhea in England between 1985 and 2000. J. Med. Microbiol. 55, 703–707; as well as unpublished data. With permission.
2006a; McLauchlin et al., 2000; Morgan et al., 2000; Muthusamy et al., 2006; Pedraza-Diaz et al., 2000, 2001b; Tiangtip and Jongwutiwes, 2002; Xiao et al., 2001a, 2002; Yagita et al., 2001), C. suis (Leoni et al., 2006a), cervine genotype (Leoni et al., 2006a; Okhuysen et al., 2002; Ong et al., 2002; TrotzWilliams et al., 2006), C. canis (Caccio, 2005; Gatei et al., 2002b; Learmonth et al., 2004; Leoni et al., 2006a), C. muris (Caccio, 2005; Gatei et al., 2002a, 2003; Guyot et al., 2001; Katsumata et al., 2000; Muthusamy et al., 2006; Palmer et al., 2003; Tiangtip and Jongwutiwes, 2002), C. andersoni (Leoni et al., 2006a), and skunk genotype (Nichols et al., 2006). The name C. pestis has been proposed for cattle strains of C. parvum because the original source was mice (Slapeta, 2006), but this name has been opposed by others owing to consideration of the International Code of Zoological Nomenclature (Xiao et al., 2007). It has been argued on the basis of evidence from the United States that genotypes from wildlife do not commonly infect humans and are of little risk to public health (Zhou et al., 2004). Although it has been suggested that C. parvum in small mammals may contribute to reinfection of sheep and cattle populations (Chalmers et al., 1997a), there is no supporting molecular evidence yet. The evidence suggests that the sources of infection for humans are predominantly human feces and the feces of agricultural animals (mostly cattle and sheep). The GP60 (also called Cpgp40/15) gene is highly polymorphic and can be used for subtyping (Table 4.2). For subtyping, seven micro- and minisatellite markers were used to examine Cryptosporidium populations from humans and animals in Scotland (Mallon et al., 2003b). They found that C. parvum isolates were grouped into three main subgroups (A–C) on the basis of strain similarities, and subtypes A and C were not detected in sheep or cattle but were in humans. Similar results have been found in other studies (Leoni et al., 2007). The results suggest either that there are human-specific isolates of C. parvum, or that we are yet to detect these genotypes in agricultural animals. With respect to the spectrum of different types, it is worth noting that the fact that some types have been found in particular animals and in individual countries does not necessarily imply that the types are restricted to this. It is likely that there is substantial adaptation by types to particular host species and the spectrum of infectivity, for a range of hosts may vary between strains.
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TABLE 4.2 Subtype Families of C. parvum and C. hominis at the GP60 (Cpgp40/15) Locus Species and Family
Source of Oocysts
Reference
C. C. C. C. C. C. C. C. C.
Human Human Human Human Human Calf, human Human, calf, sheep Human Human and calf
Trotz-Williams et al., 2006 Cohen et al., 2006 Trotz-Williams et al., 2006; Cohen et al., 2006 Trotz-Williams et al., 2006 Alves et al., 2006 Cohen et al., 2006; Mallon et al., 2003b; Trotz-Williams et al., 2006 Alves et al., 2006; Mallon et al., 2003b Alves et al., 2006; Mallon et al., 2003b Alves et al., 2006
hominis Ia hominis Ib hominis Id hominis Ie hominis If parvum IIa parvum IIb parvum IIc parvum IId
What does all the work on typing tell us about the epidemiology of cryptosporidiosis? Typing work has been useful in identifying a number of features of Cryptosporidium that have been useful in elucidating the epidemiology of human disease. It has achieved the following: • • • • • • • • • • • • • • •
Identified that not all cryptosporidiosis is caused by the same organism (Nichols et al., 1991) as originally proposed in 1980 (Tzipori et al., 1980) Identified that more than one genotype is responsible for human disease (Peng et al., 1997) Provided evidence of distinct human transmission routes and sources (Peng et al., 1997) Demonstrated that C. parvum and C. hominis are the most common species infecting humans (McLauchlin et al., 2000; Morgan-Ryan et al., 2002; Peng et al., 1997) Shown that the distribution of species differs among countries, with C. meleagridis being more common in Peru (Cama et al., 2003; Xiao et al., 2001a) Shown that the host range for species and genotypes varies (Xiao et al., 2004a) Demonstrated that outbreaks can involve more than one species and individuals can be coinfected (McLauchlin et al., 2000; Peng et al., 1997) Shown that contaminated water can contain more than one genotype (Xiao et al., 2006) Shown differences in the descriptive epidemiology for sporadic disease between C. hominis and C. parvum (Hunter et al., 2004b) Identified that some subtypes of C. parvum may be uncommon in agricultural animals but can occur in humans (Mallon et al., 2003b) Identified a range of species in immunocompromised patients (Hunter and Nichols, 2002; Cama et al., 2003) Shown the degree of genetic variation in different species (panmictic vs clonal) (Mallon et al., 2003a) Demonstrated the potential for new transmission routes (McLauchlin et al., 2000) Identified natural infection with C. hominis in cattle (Smith et al., 2005) Identified the value of intervention in reducing disease (Sopwith et al., 2005)
There are a number of issues with the use of typing. It is slightly confusing that C. parvum has been used to refer to all isolates recovered from humans before 2002 and to a subset of all isolates that are now regarded as a separate and distinct species. The taxonomy of C. parvum is further confused because the original description of this species in a mouse is probably not the organism most commonly reported as C. parvum today (Xiao et al., 2004a). The incorrect use of the term C. pestis to describe C. parvum in domestic animals has not helped (Slapeta, 2006, 2007; Xiao et al., 2007). There has been some confusion with C. parvum genotypes that have been identified in other animal species and subsequently been renamed as other species, C. hominis that was called C. parvum genotype 2 before it was renamed and C. parvum genotypes from other animal species that have yet to be renamed. The use of the term
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genotype to describe both species and types can cause confusion. As with the setting up of any new typing scheme, the terminology and comparability of isolates can lead to confusion. Confusion also arises from the recognition of different subtypes or minor species in particular hosts (see Chapter 1). Thus, C. hominis is normally not found in agricultural animals but can occasionally be found in cattle. Similarly, some subtypes of C. parvum may have been found in humans but not other animals or vice versa. Until very large numbers of isolates have been tested, it is not appropriate or helpful to say a strain is an animal or human strain because extensive testing may well prove this to be incorrect.
VII. Seroepidemiology/Population Immunity Two types of immunity have a bearing on the epidemiology of cryptosporidiosis. The first is the immune response to infection that (generally speaking) results in resolution of infection within a week or two. The second is the sustained impact of prior infection on reexposure to oocysts. Details of these are examined in Chapter 7. Within different populations there can be radically different oocyst exposure levels that can be important in the examination of outbreaks of infection. The use of serological methods to assess population exposure gives a way of determining the burden of disease over an extended time period. Methods developed in the 1980s (Campbell and Current, 1983; Koch et al., 1985; Sterling and Arrowood, 1986; Tzipori and Campbell, 1981; Ungar et al., 1986) have been extended since. They can offer advantages over other approaches in assessing community exposure (Ong et al., 2005), although the route of transmission cannot be determined. The prospective or retrospective testing for Cryptosporidium-specific antibodies can give information about recent or lifelong exposure and can be used to assess exposure through drinking water from different sources (Frost et al., 2002, 2003, 2005a). Blood samples separated by a period of time (e.g., a year) can provide an indication of the rate of seroconversion within a population and give a good indication of prevalence. This area has recently been reviewed (Casemore, 2006). Methods for detecting antibody classes include indirect immunofluorescent antibody tests using infected lamb intestine (Tzipori and Campbell, 1981) or oocysts (Casemore, 1987) as the antigen, enzyme-linked immunosorbent assay (ELISA) using sonicated oocysts (Ungar and Nash, 1986), partially purified 17-kDa oocyst antigen (Priest et al., 1999), recombinant 27-kDa antigen (Priest et al., 1999), 41-kDa antigen (Kjos et al., 2005), recombinant thrombospondin-related adhesive protein 1 (Okhuysen et al., 2004), immunoblotting (Ungar and Nash, 1986), and multiplex bead assay using recombinant C. parvum 17- and 27-kDa antigens (Moss et al., 2004). Experimental analysis of antibodies to 15-, 17-, and 27-kDa Cryptosporidium antigens produced by human volunteers suggests that IgG, IgA, and IgM antibodies to Cryptosporidium are detected within a few days of the start of symptoms, although significant amounts appear within a couple of weeks. IgM titers reach a maximum within 4 to 8 weeks and have declined substantially by 1 year (Moss et al., 1998b). IgG levels also reach a maximum within 4 to 8 weeks and are thought to remain elevated for a longer time period but decline over years. Serum and fecal IgA are produced in response to infection, but there is no detectable fecal IgM or IgG. The response to different antigens varies between immunoglobulin types. There is evidence that people exposed to Cryptosporidium infection and exhibiting a serological response to Cryptosporidium 15-, 17-, and 27-kDa antigens are at reduced risk of infection (Frost et al., 2005b). There is also evidence that prior infection can confer protection on HIV-positive individuals (Frost et al., 2005c). Serological evidence from outbreaks in Oregon (Frost et al., 1998) and Ontario (Frost et al., 2000) suggested that visitors were at greater risk of disease than the local residents because of prior exposure. A large outbreak in Devon, United Kingdom, in a holiday resort resulted in a national increase in cases that were presumed to have resulted from travel to the infected area. It has been assumed that the antibody levels detected in seroepidemiology studies are against all the species of Cryptosporidium that can cause human disease. A recent study of the IgG antibody responses of Peruvian patients to the 17- and 27-kDa antigens from the Iowa reference strain of C. parvum found responses in people infected with C. parvum, C. felis, C. meleagridis, and four subtype families of
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C. hominis (Priest et al., 2006). This study also found higher antibody levels in people who had had a previous infection and higher titers in older compared to younger children. Evidence suggests that although antibody responses are an integral part of the immune response to Cryptosporidium infection, the cell-mediated response is more important in elimination of the infection. No population-based study of the cell-mediated immune response has been conducted to date.
VIII. Infants and Children Cryptosporidiosis cases are distributed across all age groups, but in most studies of incidence or prevalence, the 1- to 2-year-old population seems to be the most affected. Because cryptosporidiosis is common in animals within the first few weeks of life, it is reasonable to assume that the comparatively low incidence in infants under 1 probably reflects reduced exposure to oocysts as a result of breast feeding or bottle feeding as well as protection from the limited oral exploration of the world in the period prior to crawling or walking. There will also be a degree of immunity derived from maternal antibodies against Cryptosporidium in the mother’s breast milk. In developing countries, malnourished children have higher rates of infection than those without malnutrition. There is also evidence that children with cryptosporidiosis are more likely to suffer from malnutrition and to die (Hunter and Nichols, 2002).
IX. Immunocompromised Populations Medical conditions that lead to compromised immunity can allow Cryptosporidium infections to be particularly severe and fail to resolve (Hunter and Nichols, 2002). These include untreated AIDS, congenital genetic immune deficiencies such as X-linked hyperimmunoglobulin M syndrome, CD40 ligand deficiency, selective IgA and Saccharomyces opsonin deficiency, and gamma interferon deficiency. Children with leukemia can get severe cryptosporidiosis, and the infection can prove a problem in bone marrow transplant patients and organ transplant patients. In AIDS patients HAART therapy is the mainstay of treatment for cryptosporidiosis (Mottet et al., 2005).
X.
Age and Sex
In England and Wales there are a larger number of cases in 1- and 2-year-olds than in any other age group (Nichols et al., 2006) (Figure 4.1). The number of male cases exceeds the number of female cases for children up to 16 years of age and then gender distribution changes dramatically, with a much higher number of cases in females than in males. The higher number of cases in male children may reflect an artifact of ascertainment (such as women taking their male children to a general practitioner more commonly than females), or a reflection of increased exposure to sources of contamination by boys. For women of child-bearing age, the exposure to young children with diarrhea may explain the reduced male/female ratio, or it may reflect the greater likelihood of women consulting a general practitioner. In older age groups, there are also more cases in women than in men, which is thought to be a reflection of the greater longevity of women.
XI. Age and Susceptibility Within populations, the more frequent occurrence of disease in young children is likely to reflect both exposure and immunity. The common occurrence of cryptosporidiosis in young animals reflects their susceptibility to infection with a low number of oocysts and common exposure to oocysts.
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FIGURE 4.1 Cryptosporidium age and sex distribution in England and Wales 1989–2005 (n = 76,065).
XII. Differences in the Epidemiology of Cryptosporidium spp. The species infecting humans are discussed in Chapters 1 and 5 and in Section VI. The strains involved in human disease appear to differ in their epidemiological characteristics.
XIII. Risk Factors In looking for causes of cryptosporidiosis, numerous studies of outbreaks and sporadic disease have been conducted, and parasitological evidence has been obtained. The evidence can provide information on risk factors that indicate likely transmission pathways (Table 4.3) as well as the sources of contamination and possible public health breakdowns that have contributed to the infections. A case-control study of sporadic cryptosporidiosis found differences in risk factors between C. parvum and C. hominis, the former being associated with animal contact and the latter with changing diapers (Hunter et al., 2004b). The study also found negative associations with eating ice cream and raw vegetables (Hunter et al., 2004b; Roy et al., 2004).
XIV. Sources Evidence for the origin of environmental oocysts is more by implication than scientific demonstration. Because oocysts are only produced naturally within the gastrointestinal and respiratory tracts of vertebrates, the sources are thought to be feces of human, agricultural, and domestic animals. The use of genetic typing has aided, and for the most part confirmed it, but there are a small but significant number of cases that may derive from other sources. Subtyping environmentally derived oocysts and those from animals may provide evidence of possible sources of fecal contamination. Although animal strains can be derived from human as well as animal feces, it is generally thought that C. hominis infections are derived from human feces. Although experimental and natural infections of animals with C. hominis have been reported, there is no evidence that this occurs commonly.
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TABLE 4.3 Risk Factors Associated with Cryptosporidiosis Risk Factor Drinking water
Private water supplies Bottled water Ice Swimming pools
Interactive water features Recreational lakes and rivers Paddling /wading pools Direct animal contact Farm visits Pets
Pet food Fairs and shows Visits to the countryside Family spread Nurseries and schools Hospital Wards with immunocompromised patients Camping Changing diapers Sexual activity Food Food and drink
Drink Unpasteurized milk Fruit juice Raw salads Flies Mollusks Food handlers
Reference Addiss et al., 1996; Atherton et al., 1995; Bridgman et al., 1995; Corso et al., 2003; D’Antonio et al., 1985; Dworkin et al., 1996; Glaberman et al., 2002; Goldstein et al., 1996; Gutteridge and Haworth, 1994; Harrison et al., 2002; Hayes et al., 1989; Howe et al., 2002; Hoxie et al., 1997; Hunter and Syed, 2001, 2002; Joseph et al., 1991; Kuroki et al., 1996; Maguire et al., 1995; McAnulty et al., 2000; McDonald et al., 2001; Naumova et al., 2003; Rodriguez-Salinas et al., 2000; Rush et al., 1990; Smith et al., 1988; Smith et al., 1989; Willocks et al., 1998; Yamamoto et al., 2000 Duke et al., 1996; Said et al., 2003 Franco and Cantusio, 2002; Leach et al., 2000 Ravn et al., 1991 1994; 2001a; 2001b; Anon. 2000; Bell et al., 1993; Hunt et al., 1994; Insulander et al., 2005; Joce et al., 1991; Lemmon et al., 1996; Lim et al., 2004; Louie et al., 2004; MacKenzie et al., 1995; Mathieu et al., 2004; McAnulty et al., 1994; Puech et al., 2001; Sorvillo et al., 1992; Stafford et al., 2000; Sundkvist et al., 1997; Wilberschied, 1995 Jones et al., 2006; Nichols, 2006 Causer et al., 2006; Kramer et al., 1998 Nichols et al., 2006 Hunter et al., 2004b; Mahdi and Ali, 2002; Pohjola et al., 1986; Preiser et al., 2003; Rahman et al., 1985; Reif et al., 1989; Stantic-Pavlinic et al., 2003b Dawson et al., 1995; Elwin et al., 2001; Evans and Gardner, 1996; Sayers et al., 1996; Smith et al., 2004; Stefanogiannis et al., 2001 Abe et al., 2002; Chalmers et al., 2002b; Cirak and Bauer, 2004; Ederli et al., 2005; Fontanarrosa et al., 2006; Hackett and Lappin, 2003; Huber et al., 2005; Lallo and Bondan, 2006; Leoni et al., 2003; McGlade et al., 2003; Mtambo et al., 1991; PedrazaDiaz et al., 2001a, 2001b Strohmeyer et al., 2006 Anon., 1996a; Ashbolt et al., 2003; Millard et al., 1994 Goh et al., 2005; Hunter et al., 2003; Smerdon et al., 2003; Strachan et al., 2003 Ribeiro and Palmer, 1986 Cruickshank et al., 1988; Hannah and Riordan, 1988 Craven et al., 1996; el Sibaei et al., 2003; Gardner, 1994; Martino et al., 1988; Ravn et al., 1991; Squier et al., 2000 Martino et al., 1988
Anon., 1996b; Smith et al., 2004 Hunter et al., 2004b Hellard et al., 2003; Pedersen et al., 1996 Dawson, 2005; Dawson et al., 2004; Duffy and Moriarty, 2003; Nichols, 2000; Rose and Slifko, 1999 Bier, 1991; Caccio and Pozio, 2001; Cook, 2003; Dawson, 2005; Dawson et al., 2004; Deng et al., 2000; Deng and Cliver, 2000; Duffy and Moriarty, 2003; Kniel and Jenkins, 2005; Laberge et al., 1996; Nichols, 2000; Rose and Slifko, 1999; Robertson and Gjerde, 2001a Garcia et al., 2006; Millard et al., 1994 Djuretic et al., 1997; Gelletlie et al., 1997 Millard et al., 1994 Robertson et al., 2005; Robertson and Gjerde, 2001b Graczyk et al., 1999b; Graczyk et al., 1999a; Graczyk et al., 2000; Hiepe and Buchwalder, 1991; Szostakowska et al., 2004 Chalmers et al., 1997b; Collins et al., 2005a; Collins et al., 2005b; Graczyk and Schwab, 2000; Li et al., 2006; MacRae et al., 2005 Quiroz et al., 2000
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TABLE 4.3 (CONTINUED) Risk Factors Associated with Cryptosporidiosis Risk Factor Travel abroad Tourist resorts Travel away from home Cruise and other ships Weather
Reference Black, 1986; Hunter et al., 2004b; Jokipii et al., 1985; Khalakdina et al., 2003; McLauchlin et al., 2000; Roy et al., 2004; Soave and Ma, 1985; Sterling et al., 1986 Cartwright, 2003 Nichols, 2003 Moss et al., 1998a; Rooney et al., 2004b; Rooney et al., 2004a Lake et al., 2005
Note: Some risk factors are yet to be confirmed by epidemiological studies.
XV. Transmission Routes The routes through which oocysts are transmitted from feces to the mouth are diverse and reflect the main transmission routes for many intestinal pathogens. Cryptosporidium differs from other pathogens such as Salmonella in its inability to grow outside the body, and a low infectious dose required to cause infection. The main routes of transmission differ between C. parvum (Figure 4.2) and C. hominis (Figure 4.3).
FIGURE 4.2 Transmission pathways for C. parvum.
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FIGURE 4.3 Transmission pathways for C. hominis.
XVI. Drinking Water The first published drinking-water-related outbreaks of cryptosporidiosis were linked to a sewage contaminated well supplying a community in Texas (D’Antonio et al., 1985) and a contaminated surface water source in Georgia (Hayes et al., 1989) (see Chapter 11). Evidence from outbreaks varies and can be categorized using an algorithm (Blackburn et al., 2004; Tillett et al., 1998). This categorization is a relatively crude device and there is always some doubt about whether an outbreak was really caused by drinking water, especially in the absence of a well-executed case-control study. In England and Wales, there were large drinking-water-borne outbreaks in 1989 and 1990 that led to the establishment of an expert group under the chairmanship of Sir John Badenoch and subsequently by Professor Ian Bouchier. The three expert reports (Badenoch, 1990; Badenoch, 1995; Bouchier, 1998) identified a number of problems with contamination of drinking water supplies, water supply management risk factors, and how to prevent outbreaks due to public water supplies. Their recommendations remain very relevant today and have greatly influenced how water companies design and operate their water treatment. In response to an outbreak in 1995 in Devon and a failure of court proceedings to obtain a successful prosecution, the government introduced additional regulations in 1999 to assess and mitigate the risk due to public water supplies. Although drinking water outbreaks accounted for only around 8% of Cryptosporidium cases, the role of public water supplies in the majority of cases is unclear. However, a strong correlation between rainfall and non-travel-related cases of cryptosporidiosis in springtime implies that many cases could have been related to water supplies lacking sufficiently robust water treatment (Lake et al., 2005). Since the change in regulations, there has been a substantial reduction in
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cryptosporidiosis in the spring that is linked to improved water treatment (Nichols et al., 2006; Sopwith et al., 2005). (See Chapter 11 for drinking water regulations in the United States.) There have been 15 drinking-water-related Cryptosporidium outbreaks in the United States between 1991 and 2002 (Craun et al., 2006b) and 55 in the United Kingdom (Furtado et al., 1998; Nichols et al., 2006; Smith et al., 2006), with outbreaks in Japan (Kuroki et al., 1996; Yamamoto et al., 2000; Yamazaki et al., 1997), Germany (Exner and Gornik, 2004), France (Cohen et al., 2006; Dalle et al., 2003), Spain (Rodriguez-Salinas et al., 2000), and Canada (Stirling et al., 2001). The reason why the United Kingdom has had so many outbreaks relative to other countries is thought to be partly due to good laboratory and disease surveillance infrastructure. Where the organism is looked for in a systematic way in diarrheal patients and reported through surveillance, outbreaks often appear. There is an implied assumption that drinking-water-related outbreaks are common in other countries but remain undetected. To prevent drinking water outbreaks, it is important to understand the problems that can cause them. I call this critical factor identification and control (CFIC) (Nichols et al., 2005). This approach has been used since the late 19th century, and a recent review of outbreaks has identified a number of examples (Hrudy and Hrudy, 2004). In the outbreak in Oxford and Swindon (Richardson et al., 1991) the recycling of backwash water to the head of the works was identified as a practice that can increase the concentration of oocysts in source waters, and this recycling is no longer used. An outbreak in Hull was caused by bypassing individual filters at a treatment works in response to a difficulty with supplying sufficient water to the community. Slow sand filtration is generally effective in removing oocysts, although waterborne outbreaks have been associated with treatment works in which slow sand filtration was used (Northern Ireland and Hull), and operational deficiencies were identified. A number of outbreaks appear to have occurred partly as a result of oocysts passing through the filters (Hayes et al., 1989). Filter efficiency varies through the cycle of their use, and the periods immediately before and after backwashing have been highlighted as posing a greater risk of oocyst breakthrough. Likewise, suboptimal clarification and flocculation processes have been suggested as a factor (Hayes et al., 1989). Ultimately, most drinking water outbreaks are caused by contamination of source waters with animal feces or sewage. The management of catchments to reduce source water contamination is an important part of the WHO Water Safety Plan approach to reducing waterborne diseases. Outbreaks have been associated with the contamination of surface waters with sewage, and there has been at least one outbreak in which contaminated surface water appears to have contaminated groundwater (Willocks et al., 1998). This outbreak was associated with other outbreaks in which there appeared to be a link between drinking water and sewage. A water–sewage cycle may have been important in an outbreak in which the location of the treatment plant may have played a role (Yamamoto et al., 2000). Drinking water outbreaks of cryptosporidiosis have been associated with heavy rainfall and flooding (Bridgman et al., 1995; Yamamoto et al., 2000). A study of cryptosporidiosis in England and Wales demonstrated associations with heavy rainfall, particularly in the spring (Lake et al., 2005). Oocyst concentrations in river water were higher in samples after rainfall compared to other sampling periods (Hansen and Ongerth, 1991), and surface water oocyst contamination after heavy rainfall is likely to cause short periods of heavy contamination. A study of 548 U.S. outbreaks of waterborne disease found that 51% of the cases were preceded by precipitation events above the 90th percentile (P = 0.002), and 68% by events above the 80th percentile (P = 0.001) (Curriero et al., 2001). For surface-water-related outbreaks, the closest associations were seen with rainfall in the preceding month, and for groundwaterrelated outbreaks, rainfall 2 months beforehand. A study examined the incidence and distribution of 92 waterborne disease outbreaks in Canada in relation to preceding weather conditions between 1975 and 2001 to test the association between high-impact weather events and waterborne disease outbreaks (Thomas et al., 2006). Total maximum degree-days above 0˚C and accumulated rainfall percentile were associated with outbreak risk. Drought can result in changes in underground water flows, and incursion of surface water into groundwater as a result of a lowered water table. A number of Cryptosporidium outbreaks have followed a dry period that ends with heavy rain (Bridgman et al., 1995; Willocks et al., 1998). Drought can also lead to less dilution of sewage effluent and animal wastes in rivers. This may have occurred in an outbreak in which the intrusion of river water into groundwater was a hypothesized transmission route (Willocks et al., 1998).
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Groundwater contamination has caused many outbreaks (Barwick et al., 2000; Bridgman et al., 1995; Lee et al., 2002; Moore et al., 1993; Willocks et al., 1998). The route of contamination from surface water to groundwater was identified in one outbreak (Bridgman et al., 1995), whereas in another the source was less clear (Willocks et al., 1998). Both outbreaks were linked to prior heavy rainfall. One outbreak resulted from a surface water drain leading directly from a field containing livestock feces (Bridgman et al., 1995). In Northern Ireland an outbreak was associated with ingress of sewage into an aqueduct (Glaberman et al., 2002). Backflow can cause an outbreak when untreated water passes into distribution (Gutteridge and Haworth, 1994), and an outbreak in Northern Ireland was caused by ingress of wastewater from a blocked drain (Glaberman et al., 2002). An outbreak in Scotland resulted when oocyst-containing water seeped into a break-pressure tank containing water for distribution, after heavy rain (Smith et al., 1989). Turbidity has increased before some Cryptosporidium outbreaks (Badenoch, 1990; MacKenzie et al., 1994; Naumova et al., 2003; Yamamoto et al., 2000), and control of turbidity can reduce the risk of oocyst contamination of drinking water. Although the relationship between oocyst contamination and turbidity is unclear (Sacco et al., 2006), removing turbidity tends to remove oocysts (Hsu and Yeh, 2003). However, water without turbidity can still be contaminated with oocysts. Higher turbidity can make oocyst recovery more difficult (DiGiorgio et al., 2002). Particle counting can give better associations with oocysts than turbidity (Brookes et al., 2005). Weather and geology can influence drinking-water-related outbreaks, but most outbreaks in England and Wales have been linked to breakdowns in water treatment and or its operation (Bouchier, 1998).
XVII. Other Drinking Waters Small sources and private water supplies are often located on farms and in rural locations in close proximity to agricultural animals. There have been six outbreaks associated with private water supplies in England and Wales (Duke et al., 1996; Nichols et al., 2006; Said et al., 2003). Contamination of these waters is related to the security of the source, safe transmission from source to tap, and whether treatment is used. Private water supply outbreaks are poorly ascertained owing to difficulties in detecting outbreaks in small communities. Outbreak investigations in which mains drinking water is the likely source of infection may flag bottled water consumption as protective in case-control studies because people have drunk this instead of mains water (Leach et al., 2000). Although bottled water can occasionally become contaminated with oocysts, especially in developing countries (Franco and Cantusio, 2002), where water safety is suboptimal, linking an outbreak to such contamination is difficult. Ice from an ice machine, contaminated by a patient with cryptosporidiosis, caused an outbreak in a hospital ward (Ravn et al., 1991).
XVIII. Recreational Water Cryptosporidium outbreaks have been linked to swimming in lakes and swimming pools (see Chapter 12 for details). Outbreaks of cryptosporidiosis have been associated with swimming pools (Anon., 2001; Bell et al., 1993; Hunt et al., 1994; Insulander et al., 2005; Joce et al., 1991; Lemmon et al., 1996; Lim et al., 2004; Louie et al., 2004; MacKenzie et al., 1995; Mathieu et al., 2004; McAnulty et al., 1994; Puech et al., 2001; Sorvillo et al., 1992; Stafford et al., 2000; Sundkvist et al., 1997; Wilberschied, 1995; Chapter 12). The contributing factors include sewage contamination of pool water through a broken drain (Joce et al., 1991), fecal accidents (Hunt et al., 1994), defective filtration, and backwashing during pool use rather than at the end of a day. Some outbreaks were in pools in which ozone treatment had been discontinued (Anon., 2000). For many outbreaks, there was no identifiable practice or feature that was linked to human disease. Outbreaks have been associated with a recreational lake in a state park in New Jersey (Kramer et al., 1998) and a recreational water park in Illinois (Causer et al., 2006). Outbreaks have also been linked to interactive water features (combinations of water jets and pools usually located
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outside) (Jones et al., 2006; Nichols, 2006). Water treatment in these features is frequently inadequate, because it is not always designed with adequate filtration. In the United Kingdom there have been two outbreaks linked to paddling pools, one to recreational use of a river and one to contamination of a stream on a beach (Nichols et al., 2006). Camping activities can result in outbreaks caused by multiple pathogens (Smith et al., 2004) as well as Cryptosporidium alone (Anon., 1996b).
XIX. Person-to-Person Transmission Transmission within families (Ribeiro and Palmer, 1986) is common, and case-control studies usually differentiate primary from secondary family cases for this reason. In the United Kingdom cases occurring in family clusters are more likely to involve C. hominis than C. parvum. Cryptosporidium can be easily transmitted in nurseries, day care settings, and schools (Cruickshank et al., 1988; Hannah and Riordan, 1988; Teresa et al., 2006). Changing diapers was a significant risk factor for C. hominis infection in a case-control study of sporadic cryptosporidiosis in the United Kingdom (Hunter et al., 2004b). Personto-person transmission of Cryptosporidium was reported more commonly in HIV-positive men who have sex with men (MSM) (homosexual men) than in IV drug users, suggesting sexual transmission (Pedersen et al., 1996). A study of MSM in Australia identified sexual behavior as a risk factor for cryptosporidiosis (Hellard et al., 2003). HIV-positive individuals can become more severely ill with Cryptosporidium infection (Jougla et al., 1996; Kim et al., 1998; Lopez-Velez et al., 1995). Disease prevention has focused on drinking bottled or boiled water, but high-risk behaviors in this group remain common (Kim et al., 1998). Transmission of Cryptosporidium within the hospital environment has been reported (Craven et al., 1996; el Sibaei et al., 2003; Gardner, 1994; Ravn et al., 1991; Squier et al., 2000) and can be a problem in units that deal with immunocompromised patients (Martino et al., 1988).
XX. Zoonotic Transmission Cryptosporidium can be transmitted from animals to humans through direct contact. This has occurred with veterinary workers (Pohjola et al., 1986; Preiser et al., 2003; Reif et al., 1989) and other people exposed to animals (Stantic-Pavlinic et al., 2003a), particularly those whose jobs are associated with agricultural animals (Mahdi and Ali, 2002; Rahman et al., 1985) and children who visit farms (Dawson et al., 1995; Elwin et al., 2001; Evans and Gardner, 1996; Sayers et al., 1996; Smith et al., 2004; Stefanogiannis et al., 2001). Educational farms need to provide hand-washing facilities for children and adults (Dawson et al., 1995; Kiang et al., 2006; Pritchard et al., 2006). An outbreak was associated with an animal nursery in a fair in Tasmania (Ashbolt et al., 2003). Infections deriving from agricultural animals are predominantly C. parvum. An outbreak of foot-and-mouth disease in the United Kingdom in 2001 led to a reduction in people’s access to the countryside, and there was a reduction in cases of cryptosporidiosis that may have been linked to this (Goh et al., 2005; Hunter et al., 2003; Smerdon et al., 2003; Strachan et al., 2003). Cryptosporidium can be detected in cats and dogs (Abe et al., 2002; Cirak and Bauer, 2004; Ederli et al., 2005; Fontanarrosa et al., 2006; Hackett and Lappin, 2003; Huber et al., 2005; Lallo and Bondan, 2006; McGlade et al., 2003; Mtambo et al., 1991; Chapter 17), and the species present are mainly C. felis and C. canis, but occasionally C. parvum and C. meleagridis (Hajdusek et al., 2004; Zhou et al., 2004). Pet food can be contaminated with Cryptosporidium (Strohmeyer et al., 2006), which may contribute to infections in these animals.
XXI.
Food and Drink
Cryptosporidium can be transmitted through food and drink (Dawson, 2005; Dawson et al., 2004; Duffy and Moriarty, 2003; Nichols, 2000; Rose and Slifko, 1999; Chapter 10). Outbreaks are difficult to
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investigate because of difficulties in typing isolates (Caccio and Pozio, 2001) and in the detection of oocysts in food (Laberge et al., 1996). Raw ingredients may be contaminated as may the water used for irrigation (Thurston-Enriquez et al., 2002) and food processing, particularly in developing countries (Sutthikornchai et al., 2005). During production and processing, raw salad can be contaminated with water containing oocysts (Robertson et al., 2005; Robertson and Gjerde, 2001b). Dairy products have caused outbreaks when the milk was improperly pasteurized (Djuretic et al., 1997; Gelletlie et al., 1997). Two outbreaks in the United States were linked to apple cider (unfermented apple juice) (Blackburn et al., 2006; Millard et al., 1994). Contamination can result from the washing process (Garcia et al., 2006) or animal feces in orchards where fruit has fallen. An outbreak of cryptosporidiosis was associated with a food handler in a cafeteria in Washington, D.C. (Quiroz et al., 2000). Foodborne outbreaks of cryptosporidiosis may occur in tourist resorts, where they are unlikely to be detected through normal surveillance processes (Cartwright, 2003). There are methods for detecting oocysts in foods (Bier, 1991; Cook, 2003; Deng et al., 2000; Deng and Cliver, 2000; Kniel and Jenkins, 2005; Robertson and Gjerde, 2001a; Chapters 6 and 10), and the impacts of various food treatments on oocysts have been examined (Dawson et al., 2004; Kniel et al., 2003). Cryptosporidium oocysts are sensitive to hot water, and it has been suggested that this can be a suitable way of cleaning and decontaminating carcasses (Moriarty et al., 2005). UV irradiation has been suggested as a treatment for fruit juices (Hanes et al., 2002), and ozonation of this product may not be effective (Blackburn et al., 2006). Flies may act as vectors for transmission of Cryptosporidium oocysts from feces to food (Hiepe and Buchwalder, 1991). Musca domestica (the housefly) can transmit Cryptosporidium oocysts in an experimental setting (Graczyk et al., 1999a, 1999b, 2000), and oocysts have been recovered from flies in a cattle barn and around a landfill site (Szostakowska et al., 2004). The epidemiology of fly transmission is difficult (Nichols, 2005), and there is no epidemiological evidence for fly transmission of cryptosporidiosis. Mollusks have been postulated as a route of transmission of Cryptosporidium (Graczyk and Schwab, 2000) because they filter large volumes of water that is commonly contaminated with animal and human fecal waste possibly containing oocysts (Gomez-Couso et al., 2006). However, infections and outbreaks have not been reported. Cryptosporidium oocysts have been detected in mussels and oysters (Chalmers et al., 1997b; Fayer et al., 1999, 2002; Freire-Santos et al., 2001; Graczyk et al., 1999c, 2001; Li et al., 2006; MacRae et al., 2005) and can survive for at least a year in seawater and for 14 days in mollusks (Tamburrini and Pozio, 1999). Improved processing of mollusks may reduce contamination (Collins et al., 2005a; Collins et al., 2005b).
XXII. Travel Cryptosporidiosis has been associated with travel abroad from the United Kingdom, Finland, and the United States (Black, 1986; Jokipii et al., 1985; Soave and Ma, 1985; Sterling et al., 1986). In the United Kingdom, travel-related cases were more commonly caused by C. hominis than C. parvum and are more commonly found in the late summer months (McLauchlin et al., 2000). Travel abroad was identified as an independent risk factor for Cryptosporidium infections in case-control studies in the United States (Khalakdina et al., 2003; Roy et al., 2004) and for C. hominis infection but not C. parvum infection in England and Wales (Hunter et al., 2004b). There is little good analytical evidence of the risk factors that are responsible for transmission within holiday resorts, although water and food quality are probably important (Cartwright, 2003). Cryptosporidium outbreaks have occurred in hotel complexes in Majorca in 2000 (C. hominis) and 2003 (C. parvum) where swimming pool water and filters were contaminated with oocysts. These large outbreaks were associated with an increase in cases throughout England and Wales.
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XXIII. Protection of Vulnerable Populations People with an acquired immunodeficiency are particularly susceptible to cryptosporidiosis and should be advised to adopt behaviors that reduce the chances of acquiring this disease (Hunter and Nichols, 2002). This includes contact with infected children, pets and agricultural animals, risky sexual behavior, swimming, and drinking unboiled mains (tap) water.
XXIV. Exclusions on the Basis of Infection Some policies for the exclusion of infected people are desirable so that transmission of cryptosporidiosis is prevented. Infected children and adults should not use swimming pools during symptomatic cryptosporidiosis (children with diarrhea should not swim) and for a period of at least 2 weeks afterward. Infants and young children should be excluded from day nurseries if they have symptomatic cryptosporidiosis.
XXV. Geographic Distribution Several reviews have looked at the distribution of cases throughout the world (Crawford and Vermund, 1988; Fayer and Ungar, 1986; Soave and Armstrong, 1986; Chapter 1). The reported rates of Cryptosporidium infection in patients with diarrhea vary substantially. General observations are that developing countries have higher rates of infection than developed ones, that disease is more common in children than in adults, and that animal contact plays an important role in developed countries. However, a simple categorization into developed and developing countries may be simplistic. Many developed countries as well as most developing countries do not have surveillance systems to detect these infections routinely, and many of the reports are “snapshot” studies of the infection over a short time period. Even in developed countries, there can be large differences in the prevalence of cryptosporidiosis over time (Nichols et al., 2006). The higher prevalence in developing countries contributes to Cryptosporidium being recognized as an important cause of traveler’s diarrhea (Black, 1986; Jokipii et al., 1985; Okhuysen, 2001; Soave and Ma, 1985; Steffen et al., 1988; Sterling et al., 1986). However, in Germany the prevalence of cryptosporidiosis in patients with traveler’s diarrhea appears to be low (Jelinek et al., 1997). A serological study provided evidence that 13.6% of Peace Corps volunteers working in West Africa became newly IgG positive over a 2-year period (Ungar et al., 1989). In England and Wales a case-control study examined the risk factors for C. hominis and C. parvum (Hunter et al., 2004b). Travel and changing diapers were significant risk factors for C. hominis, whereas animal contact was important for C. parvum.
XXVI. Seasonality The change in distribution of cases of cryptosporidiosis over time differs according to seasons. The seasonal increases in cases are linked to increased rainfall (Naumova et al., 2005), and it is assumed that this reflects contamination of source waters for drinking water from sewage and animal waste. There may also be thermal and daylight inactivation of oocysts in animal feces in the summer (Li et al., 2005) and freezing in the winter. Cryptosporidium infections were more common in the late summer in Canada (Laupland and Church, 2005). In the United States the peak incidence is in late summer (July–August) (Dietz et al., 2000; Hlavsa et al., 2005). In New Zealand C. parvum predominated in the spring and C. hominis in the autumn (Learmonth et al., 2003). In Leipzig cases are more common in the late summer (Krause et al., 1995). A seasonal distribution in Gambian children was related to times of heavy rainfall and high humidity
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(Adegbola et al., 1994). In slum children in Brazil the rate of fecal contamination with Cryptosporidium was higher in the rainy season (March–May) than in the dry season (September–December) (Newman et al., 1993). In South Africa infections were more common in late summer (January–May) (Fripp et al., 1991; Steele et al., 1989) and during periods of high rainfall (Moodley et al., 1991). Infections in GuineaBissau are at the beginning of the rainy season or just before (May–July) (Molbak et al., 1990; Perch et al., 2001). In Korea infection was more common in spring than at other times. In Kuwait in children 75% of all cases over a 2-year period occurred between January and March (Iqbal et al., 2001). Cryptosporidiosis was more common in winter months in Spain (Clavel et al., 1996; Rodriguez-Hernandez et al., 1996), but a spring and autumn seasonality in raw sewage was correlated with human cases (Montemayor et al., 2005). The assumption that seasonality should be accepted as a normal feature of the epidemiology of cryptosporidiosis has been challenged by the ability to intervene to stop the cases. A spring and autumn seasonality in cases in the North West of England changed following identification of an outbreak, the introduction of water regulations on cryptosporidiosis, and the improved water treatment (Lake et al., 2007; Nichols et al., 2006; Semenza and Nichols, 2007; Sopwith et al., 2005). This resulted in a disappearance of the spring peak that was thought to be caused by contamination of drinking water with C. parvum from animal feces.
XXVII. Burden of Disease In many countries a significant proportion of the population has had an infection before adulthood. Because infection is short-lived, the surveillance of populations with diarrheal diseases can give some feel for the extent of disease. However, when collected as routine surveillance data, it is subject to bias resulting from variations in which individuals go to physicians when they are ill, whether the practitioner requests a fecal sample, whether laboratory practice is to screen for Cryptosporidium, and whether the result will be reported to surveillance (Nichols, 2003). Because cryptosporidiosis is an important cause of large waterborne disease outbreaks and oocysts are resistant to normal disinfection processes, it is reasonable to consider that in developed countries, where disinfection of potable water is a standard practice, much of the waterborne disease burden will be due to Cryptosporidium. The disease burden from foodborne gastrointestinal infections has been examined in the United States (Mead et al., 1999) and England and Wales (Adak et al., 2002). It has been discussed at the international level (Flint et al., 2005) and recently reviewed (Roy et al., 2006). The global burden of disease associated with water, sanitation, and hygiene is estimated to be around 4% of all deaths and 5.7% of the disease burden as measured by disability-adjusted life years (DALYs) (Pruss et al., 2002). The burden of disease due to Cryptosporidium varies substantially between and within countries. An approach to developing a national estimate of waterborne disease burden has been proposed (Messner et al., 2006). However, it suffers from the same limitations as data from intervention studies and outbreaks. In particular, the degree to which a small number of intervention studies conducted in particular sites, some with methodological biases, can be used to generalize to all areas should be subjected to scrutiny. Others suggest using epidemiological measures of severity, DALYs, quality-adjusted life years (QALYs), willingness to pay, and cost-of-illness methods rather than just the number of cases (Rice et al., 2006). A workshop on estimating disease risks provides an overview of the subject, data gaps, and recommendations (Craun and Calderon, 2006b). The impact of specific interventions on the removal of oocysts from drinking water provides some indication of the extent to which this infection is locally common and transmitted by mains drinking water (tap water). The risk factors for sporadic cryptosporidiosis were examined in Cumbria, England, over 5 years (Goh et al., 2004). The incidence of sporadic cryptosporidiosis was compared in a population of 106,000 people before and after installation of a membrane filtration plant for public water supplies to that of 59,700 residents whose public water supplies were unchanged. The study was complicated by a national outbreak of foot-and-mouth disease in livestock that was associated with a decline in sporadic human cryptosporidiosis in many regions. Membrane filtration was associated with an estimated 79%
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reduction in cases after adjustment for the foot-and-mouth disease epidemic and the water source (Goh et al., 2005). A large reduction in springtime cases of C. parvum associated with improved drinking water treatment in a large area in northwest England highlighted the extent of the disease burden from this organism and the degree to which intervention can reduce it (Lake et al., 2007; Nichols et al., 2006; Semenza and Nichols, 2007; Sopwith et al., 2005).
XXVIII. Descriptive and Analytical Investigations Outbreaks provide an important focus for identifying sources of infection and improving prevention of future outbreaks. When increasing numbers of cases appear to come from a single source, based on geographic and temporal distribution, it is valuable to set up an incident control team (ICT) or outbreak control team (OCT) to investigate. One of the first functions of this group is to pull together information on the cases, including age, geographic and temporal distribution, typing to identify an outbreak strain, and the identification of potential sources, and a possible transmission route. This descriptive epidemiology can then be used to create hypotheses on the cause of the outbreak. Additional hypothesis generation can result from information obtained from affected patients. Once there are hypotheses to test, it can be useful to conduct either a case-control or cohort study using questionnaires (Craun et al., 2006a; Craun and Calderon, 2006a). Results from these analytical studies can be used to identify the source of the outbreak, and an outbreak report should be written to disseminate the findings, which may be useful in preventing future outbreaks. Alternatively, there are estimates of the amount of waterborne disease through intervention studies (Calderon and Craun, 2006; Colford, Jr. et al., 2006).
XXIX. Sporadic Disease Studies of sporadic cryptosporidiosis have attempted to determine the contributing causes (Doria et al., 2006; Goh et al., 2004; Goh et al., 2005; Hunter et al., 2004b; McLauchlin et al., 1999; Robertson et al., 2002; Roy et al., 2004). A case-control study of 152 patients and 466 unmatched controls in North Cumbria, England, found that the risk of sporadic cryptosporidiosis was associated with the volume of unboiled tap water drunk and short visits to farms (Goh et al., 2004). Most infections were due to C. parvum, and animal feces appeared to be the principal source of infection. A case-control study was conducted in seven sites of the U.S. Foodborne Diseases Active Surveillance Network (FoodNet) involving 282 cryptosporidiosis patients and 490 controls matched for geographic location and age (Roy et al., 2004). Risk factors included international travel, contact with cattle, contact with children with diarrhea, and freshwater swimming. Eating raw vegetables was protective. Case-control studies of sporadic cryptosporidiosis in 201 cases and 795 controls were recruited for Melbourne and Adelaide as well as 134 cases and 536 controls in Adelaide with different-quality water supplies (Robertson et al., 2002). Risk factors for both cities were similar, with swimming in public pools and contact with a person with diarrhea being most important. There was no association with the consumption of tap water. A case-control study of sporadic cryptosporidiosis in the United Kingdom included 427 patients and 427 controls to compare the risks between C. parvum and C. hominis. The variables that were strongly associated with all cases of illness included travel outside the United Kingdom, contact with a person with diarrhea, and touching cattle (Hunter et al., 2004b). Eating ice cream and raw vegetables were both strongly negatively associated with illness. For C. hominis infections, the most significant risk factors were travel abroad and changing diapers of children less than 5 years of age. For C. parvum, touching farm animals was associated with illness; eating raw vegetables and tomatoes were strongly negatively associated with illness. The way people with sporadic cryptosporidiosis perceive the cause of their illness is influenced by professionals they come into contact with and information from laboratory results and the media (Doria et al., 2006). These perceptions may influence the answers to questionnaires in casecontrol studies.
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XXX. Period Prevalence Cryptosporidiosis occurrence can be described by the period prevalence (usually measured in cases per 100,000 per year). The term incidence is often used in the same context, but can also be used to describe the percentage of fecal samples that are positive for Cryptosporidium. Prevalence varies geographically and can be seen to be substantially different in people of different ages and at different times of the year (Nichols et al., 2006).
XXXI. Mixed Infections Infections with two or more pathogens can occur, particularly in people returning from developing countries. It is thought to represent exposure to sources of contamination that contain multiple pathogens such as sewage or animal waste. Mixed infections have been detected with Campylobacter, Salmonella, Shigella, and Giardia (Nichols, 1992). In studies of 2414 fecal samples containing Cryptosporidium oocysts, mixtures of C. parvum and C. hominis represented nearly 1% of all cases (Leoni et al., 2006a). In outbreaks involving contaminated water, the source of contamination can be mixed, and cases infected with a mixture of species have been found in some outbreaks (Nichols and McLauchlin, 2003).
XXXII. Human Volunteer Studies The human volunteer studies conducted in Texas have provided important information about the disease in adults (Alcantara et al., 2003; Chappell et al., 1996, 1999; Dann et al., 2000; DuPont et al., 1995; Messner et al., 2001; Moss et al., 1998b; Okhuysen et al., 1998a, 1998b, 1999, 2002, 2004; Okhuysen and Chappell, 2002; Robinson et al., 2000, 2001a, 2001b; Teunis et al., 2002a, 2002b; White et al., 2000). They have identified the dose required to cause infection (DuPont et al., 1995). Oocysts were excreted in greater numbers in symptomatic patients (Chappell et al., 1996). Effects of immunity were measured by reinfection (Okhuysen et al., 1998b). The impact of hyperimmune anti-Cryptosporidium bovine colostrum on disease and oocyst excretion was also demonstrated. The work has examined the effect of antibody responses to 27-, 17-, and 15-kDa C. parvum antigens, differences in strain virulence (Okhuysen et al., 1999; Teunis et al., 2002a), the role of IFN-gamma, the neuropeptide Substance P, transforming growth factor beta1, tumor necrosis factor alpha and interleukin beta mRNA, IL15, IL8, and IL4 on human infection (Alcantara et al., 2003; Robinson et al., 2000, 2001a, 2001b, 2003; White et al., 2000) and has examined the secretion of IgA1 and IgA2 in feces post infection (Dann et al., 2000) and the reactivity to thrombospondin-related adhesive protein of Cryptosporidium (Okhuysen et al., 2004). People with previous exposure, measured by prior disease or antibodies, are less susceptible to infection (Moss et al., 1998b) and excrete fewer oocysts (Chappell et al., 1999). Prior infection provides protection to subsequent exposure with low numbers of oocysts (Chappell et al., 1999). These studies were conducted in adults. It is not clear how easily results can be applied to infants and children. Most volunteer studies were conducted with strains of C. parvum, but a C. hominis study was also conducted (Chappell et al., 2006).
XXXIII. Oocyst Viability, Infectivity, Quality, and ID50 Numerous studies have been conducted on the viability and infectivity of oocysts. There are examples of differences between strains of C. parvum and evidence that oocyst preparation can affect viability (Chapter 20). Oocysts are sensitive to some environmental chemicals (e.g., ammonia) and survive less well at warmer temperatures, in alkaline pH and if frozen. All these can contribute to deterioration of oocyst quality, both in experimental and natural situations. Oocyst viability has been reduced by orders of magnitude through the natural thermal cycles found on farms (Li et al., 2005).
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XXXIV. Risk Assessment Risk assessment can be used for strategic assessment of overall risks (Gale, 1996; Soller, 2006) or to examine the specific risks at a single site. Site-based risk assessment forms an important part of the World Health Organization water safety plan for protecting against waterborne diseases. Microbial risk assessment (MRAs) can be used in developed (Westrell et al., 2004) and developing countries (Howard et al., 2006; Soller, 2006). They have been used in the assessment of risks from root crops (Gale, 2003, 2005) and dairy calves (Nydam and Mohammed, 2005). DALYs have been used to compare the risks from drinking waterborne oocysts with that from the bromate formation resulting from ozone treatment (Havelaar et al., 2000) and shows a tenfold benefit in the reduced risks of cryptosporidiosis from drinking contaminated water compared to the risks of renal cancer from bromate. Variability in the counts used in quantitative risk assessments can be strongly influenced by variations in the recovery efficiency of the methods for detecting oocysts in water (Medema et al., 2003) as well as the genotypes of Cryptosporidium present.
XXXV. Intervention Intervention in diarrheal diseases has been important in the response to outbreaks and also in a more strategic approach to infections such as the S. enteritidis PT4 epidemic in Europe the 1990s (in this case the intervention was vaccination and improved biosecurity). With cryptosporidiosis in the United Kingdom, the large drinking-water-related outbreaks in the 1990s, particularly the Torbay outbreak in 1995 (Harrison et al., 2002; McLauchlin et al., 1998; Patel et al., 1998) and the failure to prosecute the water company stimulated the government to introduce new legislation specifically for Cryptosporidium. Legislation involved risk assessment of all drinking water treatment works and continuous monitoring for oocysts in treated water from all sites identified as at risk. The legislation was implemented in 2000–2001, and substantial new water treatment infrastructure was built in northwest England, where spring outbreaks occurred in 8 local authorities every year but decreased dramatically after intervention (Figure 4.4). From 2001 onwards infections declined, particularly C. parvum in the spring period (Figure 4.5) (Nichols et al., 2006; Sopwith et al., 2005). Membrane filtration following a period of monitoring of sporadic cryptosporidiosis resulted in a reduction of disease (Goh et al., 2005).
XXXVI. Conclusions Cryptosporidiosis is an important cause of diarrhea and the sources, risk factors, and routes of transmission are partially understood as a result of improvements in genetic typing and molecular epidemiology. For any country, the importance of routine diagnosis and reporting of Cryptosporidium to local and national surveillance organizations cannot be overemphasized. Without diagnosis of cases, outbreaks will not be identified, and without such identification, there will be little political will to reduce disease. The challenge is to identify cost-effective interventions that can be used, together with improved understanding, to reduce the burden of illness associated with this organism. Intervention studies indicate that better regulation and drinking water treatment improvements can substantially reduce disease attributed to drinking water, but disease associated with swimming may be more difficult to tackle. The causes of travel-related infections need to be examined to determine how these can be reduced. With improving protection against cryptosporidiosis in developed countries, a higher proportion of the population will not have had an infection with any Cryptosporidium spp. and may therefore be more susceptible to infection when traveling abroad or when accidentally exposed at home.
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FIGURE 4.4 Cryptosporidiosis cases per week in North West Region of England between 1996 and 2004. (From Nichols, G.L. et al., 2006. Cryptosporidiosis: A report on the surveillance and epidemiology of Cryptosporidium infection in England and Wales. Drinking Water Directorate Contract Number DWI 70/2/201. Drinking Water Inspectorate. Ref Type, Report. With permission.)
FIGURE 4.5 Annual monthly distribution of C. parvum and C. hominis in England and Wales over 6 years from 1989 to 2005. (From Nichols, G.L. et al., 2006. Cryptosporidiosis: A report on the surveillance and epidemiology of Cryptosporidium infection in England and Wales. Drinking Water Directorate Contract Number DWI 70/2/201. Drinking Water Inspectorate. Ref Type, Report. With permission.)
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Willocks, L., Crampin, A., Milne, L., Seng, C., Susman, M., Gair, R., Moulsdale, M., Shafi, S., Wall, R., Wiggins, R., and Lightfoot, N. 1998. A large outbreak of cryptosporidiosis associated with a public water supply from a deep chalk borehole. Outbreak Investigation Team. Commun. Dis. Pub. Health. 1, 239–243. Xiao, L., Alderisio, K.A., and Jiang, J. 2006. Detection of Cryptosporidium oocysts in water: Effect of the number of samples and analytic replicates on test results. Appl. Environ. Microbiol. 72, 5942–5947. Xiao, L., Bern, C., Limor, J., Sulaiman, I., Roberts, J., Checkley, W., Cabrera, L., Gilman, R.H., and Lal, A.A. 2001a. Identification of 5 types of Cryptosporidium parasites in children in Lima, Peru. J. Infect. Dis. 183, 492–497. Xiao, L., Fayer, R., Ryan, U., and Upton, S.J. 2004a. Cryptosporidium taxonomy: Recent advances and implications for public health. Clin. Microbiol. Rev. 17, 72–97. Xiao, L., Fayer, R., Ryan, U., and Upton, S.J. 2007. Response to the newly proposed species Cryptosporidium pestis. Trends Parasitol. 23, 41 and 42. Xiao, L., Lal, A.A., and Jiang, J. 2004b. Detection and differentiation of Cryptosporidium oocysts in water by PCR-RFLP. Meth. Mol. Biol. 268, 163–176. Xiao, L., Limor, J., Bern, C., and Lal, A.A. 2001b. Tracking Cryptosporidium parvum by sequence analysis of small double-stranded RNA. Emerg. Infect. Dis. 7, 141–145. Xiao, L. and Ryan, U.M. 2004. Cryptosporidiosis: An update in molecular epidemiology. Curr. Opin. Infect. Dis. 17, 483–490. Xiao, L., Sulaiman, I.M., Ryan, U.M., Zhou, L., Atwill, E.R., Tischler, M.L., Zhang, X., Fayer, R., and 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. Yagita, K., Izumiyama, S., Tachibana, H., Masuda, G., Iseki, M., Furuya, K., Kameoka, Y., Kuroki, T., Itagaki, T., and Endo, T. 2001. Molecular characterization of Cryptosporidium isolates obtained from human and bovine infections in Japan. Parasitol. Res. 87, 950–955. Yamamoto, N., Urabe, K., Takaoka, M., Nakazawa, K., Gotoh, A., Haga, M., Fuchigami, H., Kimata, I., and Iseki, M. 2000. Outbreak of cryptosporidiosis after contamination of the public water supply in Saitama Prefecture, Japan, in 1996. Kansenshogaku Zasshi. 74, 518–526. Yamazaki, T., Sasaki, N., Takahashi, S., Satomi, A., Hashikita, G., Oki, F., Itabashi, A., Hirayama, K., and Hori, E. 1997. [Clinical features of Japanese children infected with Cryptosporidium parvum during a massive outbreak caused by contaminated water supply]. Kansenshogaku Zasshi. 71, 1031–1036. Zhou, L., Fayer, R., Trout, J.M., Ryan, U.M., Schaefer III, F.W., and Xiao, L. 2004. Genotypes of Cryptosporidium species infecting fur-bearing mammals differ from those of species infecting humans. Appl. Environ. Microbiol. 70, 7574–7577. Zhou, L., Singh, A., Jiang, J., and Xiao, L. 2003. Molecular surveillance of Cryptosporidium spp. in raw wastewater in Milwaukee: Implications for understanding outbreak occurrence and transmission dynamics. J. Clin. Microbiol. 41, 5254–5257.
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5 Molecular Epidemiology*
Lihua Xiao and Una M. Ryan
CONTENTS I. II.
Introduction .................................................................................................................................. 120 Molecular Epidemiologic Tools ................................................................................................... 120 A. Concepts for Cryptosporidium Species, Genotypes, and Subtypes............................... 120 B. Genotyping Tools ............................................................................................................ 122 C. Subtyping Tools............................................................................................................... 123 D. Molecular Tools for the Analysis of Environmental Samples ....................................... 125 E. Utility of Molecular Epidemiologic Tools...................................................................... 126 III. Population Genetics of Cryptosporidium Species ....................................................................... 127 A. Tools for Population Genetic Studies: MLT and MLST................................................ 127 B. Population Genetics of C. parvum ................................................................................. 128 C. Population Genetics of C. hominis ................................................................................. 129 D. Significance of Population Genetics in Diagnostics and Epidemiology ....................... 130 IV. Molecular Epidemiology of Animal Cryptosporidiosis .............................................................. 130 Cryptosporidium Species and Genotypes in Mammals, Birds, Reptiles, and Fish....... 130 A. B. Molecular Epidemiology of Cryptosporidium Transmission in Animals ...................... 132 V. Molecular Epidemiology of Human Cryptosporidiosis .............................................................. 133 Cryptosporidium Species and Genotypes in Humans.................................................... 133 A. B. Role of Animals in Human Cryptosporidiosis Transmission......................................... 138 C. Molecular Epidemiology of Endemic Human Cryptosporidiosis.................................. 139 D. Molecular Epidemiology of Epidemic Human Cryptosporidiosis................................. 141 E. Biological, Clinical, and Epidemiologic Differences among Cryptosporidium Species ................................................................................................ 145 VI. Cryptosporidium Contamination Source Tracking ...................................................................... 146 VII. Perspective.................................................................................................................................... 150 VIII. References .................................................................................................................................... 151 IX. Appendix: Genotyping and Subtyping Cryptosporidium Oocysts in Fecal Specimens and Water Samples .................................................................................................... 164 A. Extraction of DNA in Fecal Specimen Using the QIAamp® DNA Stool Kit............... 164 B. Extraction of Cryptosporidium DNA from Oocysts Isolated from Water Samples Using Immunomagnetic Separation and the QIAamp® DNA Mini Kit ......... 165 C. Extraction of DNA from Fecal Specimen and Water Concentrates Using the FastDNA® SPIN Soil Kit.......................................................................................... 165 D. Genotyping Cryptosporidium by PCR-RFLP Analysis of the SSU rRNA Gene.......... 165 1. Primary PCR .................................................................................................... 165 2. Secondary PCR ................................................................................................ 166 3. RFLP Analysis ................................................................................................. 166 E. Subtyping C. parvum and C. hominis by PCR-Sequencing Analysis of the GP60 Gene ...................................................................................................................... 167 * 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.
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F.
I.
1. Primary PCR .................................................................................................... 167 2. Secondary PCR ................................................................................................ 168 3. Detection of Secondary PCR Products............................................................ 168 4. DNA Sequencing ............................................................................................. 168 5. Sequence Analysis ........................................................................................... 169 Multilocus Sequence Typing of C. parvum, C. hominis, and C. meleagridis ............... 169
Introduction
Because of the ability of Cryptosporidium species to infect humans and a wide variety of animals, and because of the ubiquitous presence of Cryptosporidium oocysts in the environment, humans can acquire Cryptosporidium infections through several transmission routes (Hunter and Nichols, 2002; Chapter 4). These include direct person-to-person or animal-to-person transmission and indirect waterborne and foodborne transmission, and the parasites can be of anthroponotic or zoonotic origin. The role of each transmission route in endemic areas, however, is frequently unclear because of the expensive nature of epidemiologic investigations and the inability to differentiate Cryptosporidium species by conventional microscopy. Molecular tools have been developed to detect and differentiate Cryptosporidium at the species/genotype and subtype levels (Xiao and Ryan, 2004; Caccio, 2005). The use of these tools has made significant contributions to our understanding of the biology and epidemiology of Cryptosporidium species. This includes better knowledge of the species structure and population genetics of Cryptosporidium, the roles of various transmission routes in cryptosporidiosis epidemiology, and the significance of parasite genetics in pathogenesis and clinical presentations. These recent developments have enabled researchers to make more accurate risk assessment of environmental and drinking water contamination, and have helped health officials to better educate the public on risk factors involved in the acquisition of cryptosporidiosis in vulnerable populations.
II. A.
Molecular Epidemiologic Tools Concepts for Cryptosporidium Species, Genotypes, and Subtypes
Cryptosporidium species are named based on (1) morphometric data of oocysts, (2) genetic characteristics of the parasite, (3) natural or experimental host specificity, and (4) compliance with the ICZN speciesnaming rules (Xiao et al., 2004b). There are 16 established Cryptosporidium species (see Chapter 1). However, we are in a time of rapid data accumulation, much of it genetic. Because of the uncertainty associated with Cryptosporidium taxonomy, numerous Cryptosporidium genotypes have been described without designation of species, some of which were previously lumped into C. parvum. Cryptosporidium genotypes are named after substantial sequence differences (greater than or comparable to those between established genotypes that became species) in the small subunit (SSU) rRNA or other genes are observed and phylogenetic analysis has eliminated the possibility that the differences are due to heterogeneity between copies of the rRNA gene or intragenotypic variations (see Table 5.1 for some of the commonly used primers used in the characterization of new Cryptosporidium genotypes). Although this genotype designation scheme generally reflects significant genetic differences among Cryptosporidium isolates and tends to correlate well with biological differences whenever data are available, not all genotypes differ from each other to the same extent. Thus, some genotypes exhibit extensive nucleotide differences from congenerics, whereas others are very similar to each other because of continuous host–parasite coevolution. Identifying an isolate or group of Cryptosporidium within this genus as a genotype recognizes the incompleteness of our knowledge of the parasite while acknowledging its uniqueness. A genotype is not a taxon; it is a partial and temporary descriptor, the best we have at the present time. When more data become available, a taxon designation might be made with some assurance. Species designation for
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TABLE 5.1 Some Primers Used in the Characterization of the SSU rRNA, HSP70, and Actin Genes of Various Cryptosporidium Species Gene
Primers (5′ to 3′)
Amplicon Size (bp)
SSU rRNA
SSU-F1: AACCTGGTTGATCCTGCCAGTAGTC SSU-R1: TGATCCTTCTGCAGGTTCACCTACG
~1,750
SSU rRNA
SSU-F2: TTCTAGAGCTAATACATGCG SSU-R2: CCCATTTCCTTCGAAACAGGA
~1,325
SSU-F3: GGAAGGGTTGTATTTATTAGATAAAG SSU-R4: CTCATAAGG TGCTGAAGGAGTA HSP70
Actin
~840
HSP-F1: ATGTCTGAAGGTCCAGCTATTGGTATTGA HSP-R1: TTAGTCGACCTCTTCAACAGTTGG
~2,010
HSP-F2: TA/CTTCATG/CTGTTGGTGTATGGAGAAA HSP-R2: CAACAGTTGGACCATTAGATCC Act-F1: ATGA/GGA/TGAAGAAGA/TAA/GC/TA/TCAAGC Act-R1: AGAAG/ACAC/TTTTCTGTGT/GACAAT
~1,950
Act-F2: 5′-CAAGCA/TTTG/AGTTGTTGAT/CAA Act-R2: TTTCTGTGT/GACAATA/TG/CA/TTGG
~1,066
~1,095
Usage
Reference
Amplify the full Xiao et al., rRNA gene of most 1999a eukaryotic organisms Both sets of primers Xiao et al., are Cryptosporidium 1999a, specific, and can be 2000; used as nested PCR Jiang et al., 2005b Both sets of primers Sulaiman amplify most et al., apicomplexan 2000 parasites and are used in nested PCR Both sets of primers Sulaiman amplify most et al., apicomplexan 2002 parasites and are used in nested PCR
Note: Modified from Xiao et al., 2004b. Cryptosporidium taxonomy: Recent advances and implications for public health. Clin. Microbiol. Rev. 17, 72–97. With permission.
some of the well-characterized Cryptosporidium genotypes is useful because it helps relieve much of the confusion. Caution, however, should be taken when naming new Cryptosporidium species to avoid violation of ICZN codes and unnecessary confusion in the research and public health community. This is exemplified by the recent proposal for a new species name (Cryptosporidium pestis) for a well-established Cryptosporidium species (C. parvum), which is in violation of the ICZN rules addressing the reversal of priority (Slapeta, 2006; Xiao et al., 2007a). ICZN Article 23.9.1.2 states that prevailing usage must be maintained when “the junior synonym or homonym has been used for a particular taxon, as its presumed valid name, in at least 25 works, published by at least 10 authors in the immediately preceding 50 years and encompassing a span of not less than 10 years.” Clearly, C. parvum has been accepted as a valid species for the Cryptosporidium parasites previously known as the bovine genotype in more than 25 works by over 10 authors during a period more than 10 years (Xiao et al., 2004b). In contrast, the species Cryptosporidium saurophilum in lizards (Koudela and Modry, 1998) has been accepted as a valid species for less than 10 years, thus an earlier name Cryptosporidium varanii (Pavlasek et al., 1995) buried in the literature should now be considered valid as both species appears to refer to the same intestinal parasite of lizards (Chapter 1). The term subtypes or subgenotypes is sometimes used to describe relatively minor intragenotypic variations. This is especially common for C. parvum and C. hominis, two major human pathogens. The identification of subtypes involved is frequently needed for studying the transmission dynamics in endemic cryptosporidiosis and infection and contamination sources in epidemic cryptosporidiosis. Thus, the usage of terminology of genotypes and subtypes in the Cryptosporidium research community is different from some other research fields because of the uncertainty in Cryptosporidium taxonomy. Researchers in this area should avoid the description of new Cryptosporidium genotypes based on minor sequence differences (Atwill et al., 2004; Power et al., 2004; Santín et al., 2005), which is often justifiable in other fields. In addition, researchers should avoid the use of dated species terminology (for example, C. andersoni versus C. muris) to reduce unnecessary confusion (Trout et al., 2006).
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122 B.
Cryptosporidium and Cryptosporidiosis, Second Edition Genotyping Tools
Beginning with the development of the first PCR assay for the diagnosis of Cryptosporidium in stool specimens (Laxer et al., 1991), many PCR techniques have been described for the detection of Cryptosporidium oocysts in clinical and environmental samples (Egyed et al., 2003; Smith et al., 2006b). The use of molecular techniques in the diagnosis of cryptosporidiosis, however, became popular only after the incorporation of genotyping capabilities. Since the description of the first PCR-based tool for the differentiation of C. parvum and C. hominis (Morgan et al., 1995), many genotyping tools have been developed for the characterization of Cryptosporidium epidemiology. The PCR primers are based on various antigenic, structural, housekeeping genes and unknown genomic fragments, and include various formats of detection and differentiation, including single-round and nested PCR, random amplified polymorphic DNA PCR (RAPD-PCR), arbitrary primed PCR (AP-PCR), reverse transcription PCR (RTPCR), real-time PCR, followed by restriction fragment length polymorphism (RFLP) analysis, singlestrand conformation polymorphism (SSCP) analysis, melting curve analysis, enzyme-linked immunosorbent assay (ELISA), microarray, or DNA sequencing (Egyed et al., 2003). With few exceptions, most of these techniques can efficiently differentiate C. parvum and C. hominis in stool samples, and have played a major role in understanding the transmission of human Cryptosporidium infections (Peng et al., 1997; McLauchlin et al., 2000). Their ability to detect and differentiate other Cryptosporidium species that may infect humans is largely unknown. Most of them can probably amplify DNA from C. meleagridis, but are unlikely to amplify some of the more divergent members (such as C. canis, C. felis, C. muris, and C. andersoni) of Cryptosporidium species because of the nature of most targets used by these techniques (Sulaiman et al., 1999; Jiang and Xiao, 2003; Muthusamy et al., 2006). Therefore, the use of these first-generation genotyping tools has decreased significantly. One such target, the Cryptosporidium oocyst wall protein (COWP), is still used by many researchers as a primary or confirmative genotyping method, largely because of the robustness of the technique (Pedraza-Diaz et al., 2001b; Feltus et al., 2006; Goncalves et al., 2006; Nichols et al., 2006; Soba et al., 2006; Trotz-Williams et al., 2006). Ideally, molecular diagnostic tools for Cryptosporidium should have the ability to identify all species or genotypes, or at least be able to detect all Cryptosporidium species that infect humans. Only a few such techniques are capable of doing this, largely because data on genetic diversity at species and genotype levels are only available for a few genes, such as the SSU rRNA, 70-kDa heat shock protein (HSP70), COWP, and actin (Morgan et al., 1999c; Xiao et al., 1999a, 1999c, 2002b; Sulaiman et al., 2000, 2002). Because sequence diversity among Cryptosporidium species exists over the entire HSP70, COWP, and actin genes, it is difficult to design efficient genus-specific PCR primers based on these genes, which has considerably limited their use in molecular epidemiology. Many genus-specific PCR-RFLP techniques have been described for the differentiation of Cryptosporidium species or genotypes, all based on the SSU rRNA (Awad-el-Kariem et al., 1994; Leng et al., 1996; Kimbell et al., 1999; Xiao et al., 1999c; Lowery et al., 2000; Sturbaum et al., 2001; Nichols et al., 2003; Coupe et al., 2005). The PCR method by Johnson et al. (Johnson et al., 1995) apparently can amplify various Cryptosporidium species, which can be differentiated by DNA sequencing. The SSU rRNA gene has some advantages over other genes because of the higher copy numbers and the presence of conserved regions interspersed with highly polymorphic regions, which facilitates the design of PCR primers. Nevertheless, care should be taken in choosing primer sequences because sequences conserved among Cryptosporidium species are frequently conserved among eukaryotic organisms, leading to poor specificity. This is apparently the reason for nonspecific amplification of other apicomplexan parasites by several earlier methods (Awad-el-Kariem et al., 1994; Leng et al., 1996; Kimbell et al., 1999). One disadvantage of the SSU rRNA gene is that there are minor sequence differences among different copies of the gene, which sometimes can lead to variation in RFLP for certain Cryptosporidium species or genotypes (Xiao et al., 1999b; Gibbons-Matthews and Prescott, 2003). In such situations, it is very important to differentiate intragenotypic variations among isolates from sequence variations between different copies of the gene (Atwill et al., 2004). One genus-specific genotyping tool based on ITS-1 has been described (Morgan et al., 1999b). One SSCP method based on ITS-2 has been used by one research group in genotyping Cryptosporidium
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(Gasser et al., 2003, 2004; Schindler et al., 2005). Due to extensive sequence differences between different copies of the ITS-1 or ITS-2 within a single isolate, only limited Cryptosporidium species and genotypes have been characterized at these two loci (Le Blancq et al., 1997; Gibbons-Matthews and Prescott, 2003). No matter which PCR tool is used in genotyping Cryptosporidium, all genus-specific tools have the problem with detecting only the dominant genotype in the specimen because of the inherent nature of exponential amplification by PCR and the requirement of a substantial amount of PCR product to be visible on an agarose gel. Thus, concurrent infection with mixed Cryptosporidium genotypes is more challenging to diagnose, and minor populations of species or genotypes in specimens are frequently underdetected by genus-specific tools (Reed et al., 2002). More accurate detection of the minor genotype in mixed infections would require the use of genotype-specific primers, either in combination with the use of other genotype-specific primers (Tanriverdi et al., 2003) or genus-specific primers (Cama et al., 2006a). The Cryptosporidium genotyping tool based on PCR-RFLP analysis of the SSU rRNA gene currently used in the molecular epidemiology laboratory of the Division of Parasitic Diseases, Centers for Disease Control and Prevention, is shown in the appendix.
C.
Subtyping Tools
Genotyping tools may be able to distinguish anthroponotic parasites from zoonotic parasites, but their use in epidemiologic investigation is limited by the low resolution power of these loci. As a result, subtyping tools are increasingly used in epidemiologic studies of C. parvum and C. hominis. Several types of genetic targets are used in the development of subtyping tools, including microsatellites and minisatellites (Aiello et al., 1999; Caccio et al., 2000, 2001; Feng et al., 2000; Sulaiman et al., 2001; Alves et al., 2003a; Mallon et al., 2003a; Widmer et al., 2004; Tanriverdi et al., 2006; Tanriverdi and Widmer, 2006; Gatei et al., 2007), the 60-kDa glycoprotein (GP60) gene (Strong and Nelson, 2000; Peng et al., 2001; Sulaiman et al., 2001; Leav et al., 2002; Alves et al., 2003b), double-stranded (ds) RNA element (Xiao et al., 2001b; Leoni et al., 2003a, 2003b, 2006b), and the internal transcribed spacer2 (ITS-2) of the rRNA gene (Gasser et al., 2003, 2004; Chalmers et al., 2005a; Schindler et al., 2005). These targets are used because of their higher evolutionary rates than targets used for genotyping. Microsatellites and minisatellites are DNA sequences that consist of tandemly repeated sequence motifs of 1–4 base pair (microsatellites) or more (minisatellites). Because of replication errors, these sequences generally evolve at higher rates than other nuclear genes. The genetic variations in microsatellites and minisatellites are generally variations in the number of tandem repeats, and so subtyping can be done more economically by fractionation of the PCR products, either using the conventional polyacrylamide gel electrophoresis (Feng et al., 2000; Tanriverdi et al., 2006; Tanriverdi and Widmer, 2006) or the modern GeneScan technology (Alves et al., 2003a; Mallon et al., 2003a, 2003b). However, many of the polymorphisms in microsatellites and minisatellites are in the form of single nucleotide polymorphisms (SNPs) rather than variations in the number of the tandem repeats (Sulaiman et al., 2001; Peng et al., 2003a; Gatei et al., 2006a, 2007). Thus, DNA sequencing of PCR products is widely used in the analysis of these targets (Aiello et al., 1999; Caccio et al., 2000, 2001; Enemark et al., 2002; Goncalves et al., 2006; Gatei et al., 2007). These targets are more often used in population genetic studies (see the following text). The GP60 target is similar to a microsatellite sequence by having tandem repeats of the serine-coding trinucleotide TCA/TCG/TCT at the 5′ end of the gene. However, in addition to variations in the number of trinucleotide repeats, there are extensive sequence differences in the nonrepeat regions, which categorize C. parvum and C. hominis each to several subtype families (alleles). Some of the common subtype families are Ia, Ib, Id, Ie, and If for C. hominis, and IIa, IIc, IId, and IIe for C. parvum (Figure 5.1; Table 5.2). Members of different subtype families differ from each other extensively in the primary sequences. Within each subtype family, subtypes differ from each other mostly in the number of trinucleotide repeats TCA, TCG, and TCT (mostly seen in Ie). Thus, GP60 is the most polymorphic marker identified so far in the Cryptosporidium genome, and is the most widely used Cryptosporidium subtyping target (Glaberman et al., 2002; Alves et al., 2003b, 2006b; Peng et al., 2003a; Wu et al., 2003; Zhou et al., 2003; Chalmers et al., 2005a; Sulaiman et al., 2005; Abe et al., 2006; Bjorkman and Mattsson,
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Ia
68 100
71
IIe (Anthroponotic C. parvum)
100
93
Id 81
100
IIa (Zoonotic C. parvum) 100
IIb
100
100
IId (Zoonotic C. parvum)
IIc (Anthroponotic C. parvum)
100
100
If 96
100
Ie
97 100
Ib FIGURE 5.1 Phylogenetic relationship among nine major subtype families of C. hominis (Ia, Ib, Id, Ie, and If) and C. parvum (IIa, IIc, IId, and IIe) based on a neighbor-joining analysis of the GP60 gene sequences. Anthroponotic and zoonotic C. parvum subtype families are identified.
TABLE 5.2 Major GP60 Subtype Families and Representative Sequences Species C. hominis
C. parvum
Subtype Family
Dominant Trinucleotide Repeat
Ia Ib
TCA TCA, TCG
Id Ie If Ig IIa
TCA, TCG TCA, TCG, TCT TCA, TCG TCA TCA, TCG
IIb IIc
TCA TCA, TCG
IId IIe IIf IIg IIh IIi
TCA, TCG TCA, TCG TCA TCA TCA, TCG TCA
Other Repeat (R)
GenBank Assession No.
AAAA/GCGGTGGTAAGG
AF164502 (IaA23R4) AY262031 (IbA10G2), DQ665688 (IbA9G3) DQ665692 (IdA16) AY738184 (IeA11G3T3) AF440638 (IfA19G1) EF208067 (IgA24) AY262034 (IIaA15G2R1), DQ192501 (IIaA15G2R2) AF402285 (IIbA14) AF164491 (IIcA5G3a), AF164501 (IIcA5G3b), AF440636 (IIcA5G3d) AY738194 (IIdA18G1) AY382675 (IIeA12G1) AY738188 (IIfA6) AY873780 (IIgA9) AY873781 (IIhA7G4) AY873782 (IIiA10)
ACATCA
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2006; Feltus et al., 2006; Gatei et al., 2007; Muthusamy et al., 2006; Trotz-Williams et al., 2006; Thompson et al., 2007; Xiao et al., 2007b). Unlike other subtyping targets, such as ds-RNA, ITS-2, and traditional microsatellites and minisatellites, which are generally considered nonfunctional, GP60 is located on the surface of the apical region of invasive stages of the parasite and is one of the dominant targets for neutralizing antibody responses in humans (Cevallos et al., 2000; Priest et al., 2000; Strong et al., 2000; Winter et al., 2000). Thus, it is possible to link biological characteristics of the parasite and clinical presentations with the subtype family identity (Cama et al., 2007). Cryptosporidium parvum and C. hominis are both polyphyletic in the GP60 gene, as some subtype families of C. parvum are more related to C. hominis than to other subtypes of C. parvum (Figure 5.1). Subtypes of C. parvum and C. hominis at the GP60 locus are named based on their subtype family designations and the number of each type of trinucleotide repeats (Sulaiman et al., 2005). For each subtype, the name starts with the subtype family designation (Ia, Ib, Id, Ie, If, and Ig for C. hominis, and IIa, IIb, IIc, IId, IIe, IIf, IIg, IIh, and IIi for C. parvum) followed by the number of TCA (represented by the letter A), TCG (represented by the letter G), and TCT (represented by the letter T) repeats found. Thus, the name IbA10G2 indicates that the parasite belongs to C. hominis subtype family Ib and has 10 copies of the TCA repeat and 2 copies of the TCG repeat in the trinucleotide repeat region of the GP60 gene. In the IIa subtype family, a few subtypes have two copies of the ACATCA sequence right after the trinucleotide repeats, which are represented by “R2” (R1 for most subtypes). The subtype family Ia has different copies of a 15-bp repetitive sequence 5′-AAA/G ACG GTG GTA AGG- 3′ (the last copy is 13-bp) shortly after the trinucleotide repeats, which is represented by the letter “R.” Thus, R4 indicates the presence of four copies of the 13–15-bp repeat in the Ia GP60 gene. The subtype family IIc was previously known as Ic (Strong et al., 2000). It differs from other subtype families by having no variation in the number of trinucleotide repeats (all A5G3) and by having extensive sequence polymorphism at the 3′ end of the gene. Thus, subtypes in the family are differentiated from each other by the additional letters “a” to “g,” such as IIcA5G3a (reference sequence AF164491), IIcA5G3b (reference sequence AF164501), IIcA5G3c, IIcA5G3d (reference sequence AF440621), IIcA5G3e, IIcA5G3f, and IIcA5G3g. Occasionally, a few subtypes in other families have the same trinucleotide repeat sequence, but differ from each other by one or two nucleotides in the sequence after the repeat region. They are differentiated from one another by the addition of lowercase letters, such as IIdA18G1a, IIdA18G1b, IIdA18G1c, and IIdA18G1d (Sulaiman et al., 2005). Sequencing analysis of the ds-RNA and SSCP analysis of the ITS-2 are sometimes used in subtyping C. parvum and C. hominis, but much less commonly than GP60-based techniques. The ds-RNA technique probably has the highest differentiation power, but has the disadvantage of lower sensitivity because of the RT-PCR format (Xiao et al., 2001b). As a result of its extrachromosomal presence, the ds-RNA element does not coevolve with the parasite. Therefore, ds-RNA sequences do not clearly segregate C. parvum and C. hominis into to two separate groups (Xiao et al., 2001b). One SSCP method based on ITS-2 has been used by one research group in subtyping C. parvum and C. hominis (Gasser et al., 2003, 2004; Chalmers et al., 2005a; Schindler et al., 2005). It remains to be determined whether this technique is affected by the extensive sequence differences between different copies of the ITS-2 (Le Blancq et al., 1997). It is important to keep in mind that most subtyping tools were developed based on nucleotide sequences of C. parvum or C. hominis. Even though some of these tools will probably amplify DNA of Cryptosporidium species closely related to these two species, such as C. meleagridis, they usually do not amplify DNA of distant Cryptosporidium species and genotypes, such as C. canis, C. felis, C. suis, and C. muris, which can be found in humans. Thus, subtyping has largely been restricted to C. parvum and C. hominis. Nevertheless, GP60- and HSP70-based PCR tools have been used effectively in subtyping C. meleagridis (Glaberman et al., 2001).
D.
Molecular Tools for the Analysis of Environmental Samples
Diagnosis of Cryptosporidium to the species/genotype level is especially a challenge for environmental samples because of the occurrence of usually a very low number of oocysts and the rich presence of PCR inhibitors, but is essential for the assessment of public health importance and contamination sources
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of Cryptosporidium oocysts. The identification of Cryptosporidium oocysts in environmental samples is largely made by the use of immunofluorescent assays (IFAs) after concentration processes (EPA methods 1622 and 1623, and their equivalents in other countries) (Sinclair, 2000; Lindquist et al., 2001). Because IFA detects oocysts from all Cryptosporidium species, the species distribution of Cryptosporidium oocysts in environmental samples cannot be assessed (Weintraub, 2006). Although many surface water samples contain Cryptosporidium oocysts, it is unlikely that all of these oocysts are from humanpathogenic species or genotypes, because only five Cryptosporidium species (C. parvum, C. hominis, C. meleagridis, C. canis, and C. felis) are responsible for most human Cryptosporidium infections (Xiao et al., 2004b). Information on the source of Cryptosporidium contamination is necessary for effective evaluation and selection of management practices for reducing Cryptosporidium contamination in source water and determining the risk of cryptosporidiosis. Thus, identification of oocysts to the species and genotype level is of public health importance. The performance of many PCR methods in the analysis of environmental samples have been evaluated with Cryptosporidium-negative samples seeded with known numbers of C. parvum oocysts. PCR was performed on DNA extracted directly from water concentrates seeded with Cryptosporidium oocysts either without isolating the oocysts or using Percoll-sucrose flotation. Variable sensitivities were obtained, ranging from 1 to >100 oocysts per sample. Many researchers reported the inhibitory effect of surface water on PCR (Johnson et al., 1995; Rochelle et al., 1997; Sluter et al., 1997; Chung et al., 2000; Lowery et al., 2000; Xiao et al., 2000). Thus, more recent techniques have used an oocyst isolation step using immunomagnetic separation (IMS) prior to DNA extraction to remove PCR inhibitors or contaminants present in water samples (Johnson et al., 1995). The sensitivity of the IMS-PCR is up to 1,000-fold higher than when IMS was not used prior to DNA extraction (Johnson et al., 1995), and is generally higher than the EPA method 1622/1623, which is based on immunofluorescent microscopy (Xiao et al., 2000, 2006b). A strategy combining direct DNA extraction using the FastDNA soil DNA extraction kit and neutralization of residual PCR inhibitors using high concentrations of nonacetylated bovine serum albumin in PCR has been used successfully in the PCR detection of Cryptosporidium oocysts in water samples (Jiang et al., 2005a). The protocol for this is in the appendix at the end of the chapter. Earlier successful reports of PCR analysis of natural water samples were mostly done using conventional PCR methods (Johnson et al., 1995; Mayer and Palmer, 1996; Kaucner and Stinear, 1998; HallierSoulier and Guillot, 2000). Unfortunately, these techniques do not have a species differentiation/genotyping component; thus, the identity of Cryptosporidium was not established. Since then, various SSU rRNA-based PCR-RFLP, PCR-sequencing, or real-time PCR tools have provided useful data on the genotype, source, and human infection potential of Cryptosporidium oocysts in water in various areas (Xiao et al., 2000, 2004c, 2006b; Lowery et al., 2001b; Jellison et al., 2002; Sturbaum et al., 2002; Ward et al., 2002; Nichols et al., 2003; Jiang et al., 2005b; Ruecker et al., 2005, 2007; Coupe et al., 2006; Hashimoto et al., 2006; Hirata and Hashimoto, 2006; Masago et al., 2006; Nichols et al., 2006; Sunnotel et al., 2006). A commonly used SSU-based PCR-RFLP tool is shown in the appendix (Xiao et al., 2000, 2001c, 2004c, 2006b; Tsushima et al., 2003; Zhou et al., 2003; Jiang et al., 2005b; Ruecker et al., 2005, 2007; Alves et al., 2006a).
E.
Utility of Molecular Epidemiologic Tools
The development of molecular tools for genotyping and subtyping has been useful in understanding host specificity Cryptosporidium species and the transmission and clinical presentation of human cryptosporidiosis. The following are examples of the use of molecular tools in epidemiologic investigations of human infections. 1. Establishment of the identity of Cryptosporidium in humans. We can now identify the species of Cryptosporidium that infect humans, the proportion of infections attributable to each species in various socioeconomic and epidemiologic settings, the temporal and geographic variation in the occurrence of various Cryptosporidium species in humans, and the heterogeneity within each species causing human infection.
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2. Identification of infection or contamination sources. When used in conjunction with traditional epidemiologic and environmental investigations, molecular tools can help identify the source of infection or contamination: anthroponotic versus zoonotic Cryptosporidium infection, and farm animal or companion animal origin versus wildlife origin. With a large sample size, molecular tools can help assess the human infective potential of Cryptosporidium species and genotypes from various animals that are in frequent contact with humans. With higher-resolution tools, molecular techniques can make a direct linkage between human cases of cryptosporidiosis and contamination sources (contaminated food item or water source, human index case such as a food handler, animal reservoir, etc.). 3. Characterization of the transmission dynamics of cryptosporidiosis in communities. Highresolution molecular tools can help distinguish cryptosporidiosis point-source outbreaks from endemic but unrelated clusters of cases. These tools may serve to assess the intensity (high versus low complexity of parasites in the community) and nature of the endemic (stable transmission versus new introduction of parasites), identify common transmission pathways, distinguish multiple episodes of infections in humans, elucidate mechanisms of immunity against homologous and heterologous Cryptosporidium species, and differentiate new episodes of infection from reactivation of latent infections. 4. Characterization of the clinical spectrum and pathobiology of cryptosporidiosis. Molecular tools can improve understanding of the mechanisms underlying the variable clinical presentations and attack rates in outbreaks, variation in disease spectrum in the general public and AIDS patients, and differences in infection sites and pathophysiology of Cryptosporidium species and genotypes. In addition to host susceptibility, it is likely that the genetic diversity of Cryptosporidium species plays an important role in the variation in clinical and pathologic spectrum of human cryptosporidiosis
III. Population Genetics of Cryptosporidium Species With the development of Cryptosporidium subtyping tools, it has become possible to assess the genetic and population structure of Cryptosporidium species. Earlier studies concentrated on the presence or absence of recombination between C. parvum and C. hominis, using multilocus sequence analysis of conserved gene targets. Consistent with other apicomplexan parasites, despite the presence of a sexual phase in the life cycle, common occurrence of coinfection, and an overlap in infection site, there is no evidence for genetic recombination between the two common Cryptosporidium species C. hominis and C. parvum (Morgan-Ryan et al., 2002; Widmer, 2004). The only noticeable exception is the polyphyletic nature of the two species in the GP60 gene, with mosaic sequences observed between some subtype families of C. parvum and C. hominis (Leav et al., 2002). Even though this has been suggested as an indicator of recombination between the two species (Leav et al., 2002), results of multilocus sequencing suggest that evolution of the GP60 subtype families might have preceded the species divergence (Gatei et al., 2006a). Earlier studies of the intraspecific population structure of Cryptosporidium used mostly genetic targets that were not highly polymorphic (Glaberman et al., 2001; Sulaiman et al., 2001). Highresolution multilocus typing tools are now used to characterize the genetics and population structure of C. parvum and C. hominis and the role of host, geographical, and temporal factors in the evolution of these two species (Mallon et al., 2003a; Ngouanesavanh et al., 2006; Tanriverdi et al., 2006).
A.
Tools for Population Genetic Studies: MLT and MLST
The recent genome sequencing of C. parvum and C. hominis (Abrahamsen et al., 2004; Xu et al., 2004) has provided the basis for the development of high-resolution typing tools for the characterization of these two species. Thus far, the complete genomes of C. parvum Iowa isolate and C. hominis TU502 isolate, and the chromosome 6 of C. parvum Iowa isolate maintained in a different laboratory have been sequenced. This has allowed the identification of microsatellite and minisatellite sequences in C. parvum
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and C. hominis genomes and other targets highly polymorphic between C. parvum and C. hominis or between the C. parvum Iowa isolate maintained in different laboratories (Cama et al., 2006b). Two categories of typing techniques are used to determine population structure of C. parvum and C. hominis. In multilocus typing (MLT), variation in microsatellites and minisatellites are assessed on the basis of length variations using polyacrylamide gel electrophoresis (Feng et al., 2000; Tanriverdi et al., 2006; Tanriverdi and Widmer, 2006) or the GeneScan technology (Alves et al., 2003a; Mallon et al., 2003a, 2003b; Ngouanesavanh et al., 2006). This allows the use of many targets in the MLT techniques economically. For example, using 7 minisatellite and microsatellite markers, 38 multilocus subtypes were identified in 180 C. parvum and C. hominis isolates (Mallon et al., 2003a) and 48 multilocus subtypes in 240 C. parvum isolates in Scotland (Mallon et al., 2003a, 2003b). In contrast, 14 markers identified 14 multilocus subtypes in 61 C. parvum isolates from Israel (Tanriverdi et al., 2006). The second category of typing techniques, multilocus sequence typing (MLST), relies on the detection of genetic heterogeneity by DNA sequencing of the amplified PCR products (Glaberman et al., 2001; Sulaiman et al., 2001; Gatei et al., 2007). Comparing to PCR product fractionation, MLST allows more definitive detection of sequence polymorphism and the inclusion of makers with single-nucleotide polymorphisms (SNPs) (Sulaiman et al., 2001; Gatei et al., 2007). As shown previously with the HSP70 locus (a minisatellite marker used in the Scottish MLT technique), significant variation in minisatellite loci is in the form of SNPs, which would have been missed by typing techniques based on length polymorphism (Glaberman et al., 2001; Sulaiman et al., 2001; Peng et al., 2003a). This is seen in other minisatellite markers such as Mucin1 and MSC6-7 (Gatei et al., 2007). Even for the highly polymorphic microsatellite locus GP60, many subtypes in each family have the same length even though the sequences are different. For example, C. hominis subtype family Id has two common subtypes in children in Kolkata, IdA15G1 and IdA16, which had the same length of PCR products despite the sequence difference in the microsatellite repeat region (Gatei et al., 2007). Similarly, many common C. hominis (such as IbA10G2 and IbA9G3) and C. parvum (such as IIaA15G2R1 and IIaA16G1R1) subtypes have the same length of the GP60 gene, and the anthroponotic C. parvum IIc subtypes have no variation in the number of microsatellite repeats. The more expensive nature of MLST has limited the number of loci targeted and the size of specimens under analysis. For example, seven microsatellite, minisatellite, and SNP markers identified 25 multilocus sequence types in 37 isolates of C. hominis from children in Kolkata, India (Gatei et al., 2007). Because studies of linkage disequilibrium (LD) usually examine the association of subtypes among loci on the same chromosome, most of the markers used in our laboratory are located in chromosome 6 (Gatei et al., 2007).
B.
Population Genetics of C. parvum
Of C. parvum populations infecting humans and livestock in Aberdeenshire, United Kingdom, LD analysis between pairs of loci combined with measures of genetic distance and similarity demonstrated one large population of C. parvum comprising both human and bovine isolates. This group of C. parvum had a panmictic population structure and was in linkage equilibrium, suggesting that genetic exchange occurred frequently (Mallon et al., 2003a). Thus, new subtypes can emerge as a result of sexual recombination during concurrent infection of two C. parvum subtypes, which was demonstrated previously by experimental infection of mice with different subtypes of C. parvum (Feng et al., 2002). The same large population of C. parvum in both humans and cattle was seen in several areas in Scotland (Mallon et al., 2003b). In contrast, bovine C. parvum in Israel has a clonal population structure (Tanriverdi et al., 2006). Significant LD was observed between 14 microsatellite and minisatellite markers in 61 isolates from diverse areas. Significant geographic segregation in C. parvum subtypes was observed in both Israel and Turkey, as distinct multilocus types were often detected in individual herds. It was concluded that even in a clonal population structure, genetically distinct populations of C. parvum can emerge within a group of hosts in a relatively short time (Tanriverdi et al., 2006). In Scotland, however, there is seemingly no geographical or temporal substructuring within C. parvum (Mallon et al., 2003b). In a recent multilocus characterization of C. parvum from humans and animals in France and Haiti, two major populations of C. parvum were identified, with one population showing panmixia and one
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showing epidemic population structure (Ngouanesavanh et al., 2006). Whether the finding of two major groups of C. parvum GP60 subtypes in humans in Kuwait City represents an epidemic population remains to be decided using MLT or MLST tools (Sulaiman et al., 2005). In the Scottish studies, two C. parvum populations were found only in humans (Mallon et al., 2003a, 2003b). The presence of human-adapted C. parvum subtypes is well known at the GP60 locus, as the anthroponotic subtype families IIc and IIe (Figure 5.1) have been found in humans in South Africa, Uganda, Portugal, the United States, and Peru, but not in animals (Leav et al., 2002; Alves et al., 2003b, 2006b; Xiao et al., 2004b; Xiao and Ryan, 2004; Akiyoshi et al., 2006). In some of the areas, the zoonotic IIa subtype family is rarely seen in humans. Tanriverdi and associates theorized that humans and cattle in the same geographic areas harbor distinct C. parvum populations, and such host substructuring is the driving force for the emergence of host-adapted Cryptosporidium species and genotypes (Tanriverdi et al., 2006). This is supported by findings in France and Haiti (Ngouanesavanh et al., 2006). It is important to note that these human-adapted C. parvum subtypes are not the various host-adapted Cryptosporidium genotypes previously described based on sequence analysis of conservative genes such as SSU rRNA, actin, and HSP70, as the former would have minimal sequence variations at these loci and the latter emerged millions of years ago (Xiao et al., 2002b). The difference in the population genetic structure of C. parvum from different areas awaits further validation but has received support from findings of C. parvum human isolates having a clonal population structure in France and an epidemic population structure in Haiti (Ngouanesavanh et al., 2006). Other protozoa have a considerable variation within specific parasites, with substructures ranging from clonal, epidemic, to panmictic that may depend on diverse factors including transmission dynamics, geographical regions, and host-specificity (Tibayrenc and Ayala, 2002; Lehmann et al., 2004; Oura et al., 2005). Thus, it is possible to have a clonal population of C. parvum in one area but a panmictic population in another. There is evidence that in one area, C. parvum can have an epidemic population structure in humans but a panmictic population structure in cattle (Mallon et al., 2003b), or a clonal population structure in humans but an epidemic population structure in animals (Ngouanesavanh et al., 2006). Data from more epidemiologic settings using more polymorphic loci are needed before firm conclusions on the population structure of C. parvum can be made (Tait et al., 2004; Widmer, 2004).
C.
Population Genetics of C. hominis
Although complete LD was shown in an initial population genetic study of C. hominis (Sulaiman et al., 2001), many specimens were from outbreaks in the United States, and the markers used were mostly not very polymorphic. In Scotland, C. hominis displayed a lower number of subtypes than C. parvum (Mallon et al., 2003a). No differences in heterogeneity, however, were observed between C. parvum and C. hominis in a small number of isolates in the United States (Tanriverdi and Widmer, 2006). In Scotland, because most isolates belonged to only two major multilocus subtypes (89% of the isolates), it was concluded that the population structure of C. hominis was clonal. However, these two multilocus subtypes differed from each other only at one locus (MS5), which made it impossible to calculate LD (Mallon et al., 2003a). The Scottish specimens were from a C. parvum-endemic area; thus, it was not clear if the observed low number of subtypes in C. hominis was due to the epidemic population structure of the parasite (Ngouanesavanh et al., 2006), point source infections, the limited number of specimens, or the level of resolution of the genetic markers. Nor was it possible to assess if differences in human infectivity among C. hominis subtypes could lead to unique population substructures compared to other species of Cryptosporidium. Low genetic diversity was seen in MLT of C. hominis from France and Haiti, which was interpreted as evidence for an epidemic population genetic structure (Ngouanesavanh et al., 2006). MLST analysis of 37 C. hominis isolates from children in Kolkata, India (Gatei et al., 2007), found 25 multilocus subtypes by combined sequence length polymorphism and SNP analysis, which formed four distinct groups in this population. The strong and significant LD between sites based on the multilocus gene sequence and the lack of variation at the normally polymorphic HSP70 locus suggest the population structure was clonal. In addition, while structuring was extensive (25 multilocus subtypes in 37 specimens), predominant subtypes were observed at all loci. This observation was supported by the significant Zns and Fs values that suggest molecular selection. Selection would explain the previously
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observed reduced subtype heterogeneity in C. hominis compared with C. parvum. This selection does not appear to be due to an expansion of a single subtype as seen in the isolates analyzed in Scotland (Mallon et al., 2003a) and as would be the case in an epidemic clonal structure. Furthermore, rapid demographic expansion in a population usually gives large negative Fs values, which was not seen. The absence of significant recombination and the strong intragenic LD are indicative of a stable clonal endemicity in this population. The formation of four groups in the phylogeny based on the Bayesian method, however, was probably due to extensive sequence differences among C. hominis subtype families at the GP60 locus. The significance of the diverse MLST within C. hominis in relation to geographical and temporal factors and clinical manifestations of disease has not been investigated.
D.
Significance of Population Genetics in Diagnostics and Epidemiology
The Cryptosporidium genetic structure has direct implications for understanding its biology as well as characterizing transmission routes and dynamics and infection sources in specific areas. A better understanding of the parasite population structure has had a profound impact on our knowledge of likely transmission routes and infection sources of the parasite in the community as demonstrated by the finding of (1) human-adapted C. parvum populations (Mallon et al., 2003a, 2003b), (2) distinct multilocus subtypes between humans and cattle in the same area (Tanriverdi et al., 2006), and (3) different C. parvum population structures between humans and cattle in one area (Mallon et al., 2003b; Ngouanesavanh et al., 2006). It is very likely that anthroponotic transmission of C. parvum may be more common than previously believed, not only in developing countries but also in C. parvum endemic areas. The finding of geographically distinct subtypes of C. parvum has significant implications to understanding the transmission dynamics of cryptosporidiosis in different areas (Gatei et al., 2006a). In Israel and Turkey, most cattle farms had distinct C. parvum subtypes, which probably resulted from the rapid emergence of novel subtypes within a group of structured hosts (Tanriverdi et al., 2006). Thus, farm management practices such as animal free ranging and frequent animal trade increase C. parvum heterogeneity on farms and the complexity of infection (occurrence of mixed subtypes) (Tanriverdi et al., 2006). The characterization of the population genetic structure of Cryptosporidium species is necessary in high-resolution tracking of infection or contamination sources and in outbreak investigations. A clonal population structure indicates that multilocus subtypes are relatively stable in time and place, and thus can be used effectively in the longitudinal tracking of the transmission in the community and in investigation of outbreaks. A panmictic population structure suggests that multilocus subtypes may not be stable in time and place, which might present a problem in temporal tracking of transmission but allow spatial tracking (Mallon et al., 2003a). An epidemic population structure is indicative of recent introduction of the parasite population into the community and an unstable epidemic of the disease. The power of MLST in the genetic comparison of the C. parvum Iowa isolate maintained in different laboratories demonstrated that oocysts used in sequencing the complete C. parvum genome were not from the original C. parvum Iowa isolate (Cama et al., 2006b).
IV. A.
Molecular Epidemiology of Animal Cryptosporidiosis Cryptosporidium Species and Genotypes in Mammals, Birds, Reptiles, and Fish
There is extensive genetic variation within the genus Cryptosporidium (Figure 5.2). In addition to the 16 accepted species of Cryptosporidium, nearly 40 Cryptosporidium genotypes have been described and new genotypes are continually being discovered (Xiao et al., 2004b; Chapter 1, this book). Phylogenetically, Cryptosporidium species and genotypes form two groups: those found primarily in the intestine, and others in the stomach. Each group contains parasites of mammals, birds, and reptiles. The placement of fish parasites, however, is not clear. Most species and genotypes are host-adapted in nature, having a narrow spectrum of natural hosts. The biological and taxonomic significance of most genotypes has been reviewed (Xiao et al., 2004b; Chapter 1, this book). In recent years, several well-known genotypes
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C. wrairi 517 (8) Ferret genotype 351 (4) C. parvum 6 (52) Mouse genotype 411 (20) Skunk genotype (10) 99 Opossumgenotype I 1041 (8) Marsupial genotype I428 (9) Horse genotype 6481 (1) Rabbit genotype 2244 (2) 83 C. hominis monkey genotype 44 (3) 70C. hominis 120 (122) C. meleagridis 295 (18) C. suis 427 (8) Marsupial genotype II 6475 (2) Cervine genotype 6482 (13) Fox genotype 2041 (1) Muskrat genotype II 3657 (6) 70 Deer mice genotype 1457 (2) Opossum genotype II 1040 (12) Squirrel genotype 7406 (15) Muskrat genotype I 5490 (30) Bear genotype 961 (1) C. canis coyote genotype 2011 (2) 60 98 98 C. canis dog genotype 244 (15) 92 C. canis fox genotype 5035 (11) C. felis 288 (10) C. varanii 340 (15) 98 Goose genotype I 886 (36) 89 Goose genotype II 1182 (11) Duck genotype 6876 (2) 99 Pig genotype II 6734 (2) C. bovis 2622 (30) 77 92 Deer genotype 2040 (8) 81 Deer-like genotype 6293 (15) 100 78 Snake genotype 938 (1) C. baileyi 39 (6) Tortoise genotype 750 (5) 96 C. serpentis 63 (65) Lizard genotype 1665 (1) 93 95 C. andersoni 20 (18) C. muris 34 (15) Woodcock genotype 5391 (1) 100 C. galli finch genotype 1436 (2) C. galli 4300 (7) Eimeria tenella AF026388
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FIGURE 5.2 Genetic relationship among named Cryptosporidium species and genotypes inferred by a neighbor-joining analysis of the partial SSU rRNA gene. Values on branches are percent bootstrapping using 1000 replicates. Numbers following species or genotypes are isolate identifications used in the construction of the phylogenetic tree, whereas numbers in parentheses are number of isolates sequenced. The two established Cryptosporidium spp. in fish are not included as they have not been characterized genetically (based on Xiao et al., 2004b. Cryptosporidium taxonomy: Recent advances and implications for public health. Clin. Microbiol. Rev. 17, 72–97. With permission).
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have been elevated to species status as sufficient biologic data become available. Cryptosporidium hominis was previously known as C. parvum genotype I or human genotype, C. suis was previously known as pig genotype I, and C. bovis was previously known as bovine genotype B (Morgan-Ryan et al., 2002; Ryan et al., 2004; Fayer et al., 2005). This trend is likely to continue. The host-adaptation and host-parasite coevolution is a continuous process. Thus, even though a Cryptosporidium genotype in Australian marsupials, marsupial genotype I, is genetically related to opossum genotype I found in the American marsupial, there are substantial genetic differences between the two Cryptosporidium genotypes (Xiao et al., 2002b). Likewise, C. canis found in dogs, coyotes, and foxes in the United States differed from each other slightly in the SSU rRNA gene sequences, even though the divergence of these animal species was fairly recent (Xiao et al., 2002b). The same is true for C. bovis isolates from cattle and yaks (Feng et al., 2007). Because of the close relatedness of these newly divergent genotypes, occasionally, cross-species transmission can occur, such as finding C. canis dog genotype in a fox (Zhou et al., 2004a), the Cryptosporidium duck genotype in a Canada goose (Zhou et al., 2004b), and the C. hominis monkey genotype in two humans (Mallon et al., 2003a).
B.
Molecular Epidemiology of Cryptosporidium Transmission in Animals
The existence of host-adapted Cryptosporidium species or genotypes indicates that cross transmission of Cryptosporidium among different groups of animals is probably limited. Surveys conducted in pigs, kangaroos, squirrels, fur-bearing mammals, Canada geese, and reptiles have shown that most animals are infected with only a few host-adapted Cryptosporidium species or genotypes (Guselle et al., 2003; Jellison et al., 2004; Power et al., 2004; Xiao et al., 2004d; Zhou et al., 2004a, 2004b). The transmission dynamics of C. parvum are more complicated. It was once thought that C. parvum infected nearly all mammals, with widespread cross-species transmission (Tzipori and Griffiths, 1998). This was largely due to reliance on microscopy of oocysts, which suffered from the lack of distinguishable morphologic differences among most Cryptosporidium species and genotypes. It is now generally accepted that C. parvum (previously referred to as the bovine genotype) primarily infects ruminants and humans, even though natural infections have been found occasionally in other animals such as mice and raccoon dogs (Morgan et al., 1999d; Matsubayashi et al., 2004). Although C. parvum was detected in a few horses, its prevalence is not known (Grinberg et al., 2003; Hajdusek et al., 2004; Chalmers et al., 2005b), and horses are known to be infected with a Cryptosporidium horse genotype (Ryan et al., 2003). Even within ruminants, C. parvum seems to be mostly a parasite of cattle. In Australia and the United States, C. parvum infection appears not very common in sheep, which are more often infected with the Cryptosporidium cervine genotype and other genotypes (Ryan et al., 2005b, Santín et al., 2007a, 2007b). Few Cryptosporidium species infect a broad range of hosts. A noticeable exception is the Cryptosporidium cervine genotype. Since its initial finding in storm water, it has been found in domestic and wild ruminants (sheep, mouflon sheep, blesbok, nyala, and deer), rodents (squirrels, chipmunks, woodchucks, raccoons, and deer mice), and primates (lemurs and humans) (Xiao et al., 2000; Perz and Le Blancq, 2001; Ong et al., 2002; da Silva et al., 2003; Ryan et al., 2003, 2005b; Blackburn et al., 2006; Feltus et al., 2006; Leoni et al., 2006a; Soba et al., 2006; Trotz-Williams et al., 2006). Because it is the most common Cryptosporidium found in pristine water, it is likely some other wild mammals are also hosts (Jiang et al., 2005b; Xiao et al., 2006b). In cattle, there is an age-associated occurrence of different Cryptosporidium species and genotypes. Most Cryptosporidium infections in preweaned calves are due to C. parvum, whereas most Cryptosporidium infections in postweaned calves are due to C. bovis and the deer-like genotype, which can be frequently found in yearlings and adult cattle. Cryptosporidium andersoni is first found in juveniles, but more frequently in yearlings and adults (Santín et al., 2004; Fayer et al., 2006; Feng et al., 2007; Langkjaer et al., 2007; Chapter 18, this book). Cryptosporidium bovis and Cryptosporidium deer-like genotypes are genetically related, and the age pattern of the host is very similar for both parasites, even though the deer-like genotype is usually less common. Because both are sometimes seen in preweaned calves in the absence of C. parvum infection, and because the oocyst shedding intensity of the two parasites is several logs lower than that of C. parvum, it is not known if C. bovis and the deer-like genotype infect young cattle, but they are obscured by overwhelming numbers of C. parvum oocysts.
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Age-associated infection with Cryptosporidium species and genotypes has not been well documented for other animal species. However, C. suis appears more common in preweaned piglets, and pig genotype II more common in older pigs (Langkjaer et al., 2007). It would not be surprising to see the same phenomenon in other animals such as sheep and deer. Geographic differences in the diversity of C. parvum in cattle and distribution of subtypes have been reported based on GP60 subtyping (Xiao et al., 2007b). In the United States, Canada, and the United Kingdom, calves were infected with only the IIa subtype family of C. parvum (Peng et al., 2003b; Thompson et al., 2007; Trotz-Williams et al., 2006; Xiao et al., 2007b). In Portugal, Italy, and Serbia, although IIa subtypes were dominant, IId subtypes were found occasionally (Alves et al., 2003b, 2006b; Wu et al., 2003; Misic and Abe, 2007). Subtype diversity of C. parvum in cattle is very high in Northern Ireland and Serbia, modest in Canada and Michigan, and very low in Portugal and along the East Coast of the United States (Alves et al., 2003b, 2006b; Peng et al., 2003b; Misic and Abe, 2007; Thompson et al., 2007; Trotz-Williams et al., 2006; Xiao et al., 2007b). GP60 sequence analysis of 34 bovine samples from 12 farms in Michigan revealed 6 subtypes from the family IIa. Many farms had more than one subtype, suggesting that, even under intense transmission in farm animals, heterogeneous Cryptosporidium were circulating in a broad geographic area and on specific farms (Peng et al., 2003b). Similar results were found in neighboring Ontario Province, Canada (Trotz-Williams et al., 2006). In contrast, only six IIa subtypes were seen in calves in eight states in eastern United States, and many states had only one subtype (Xiao et al., 2007b). Subtype IIa15G2R1 is the most widely distributed IIa subtype, found in cattle in the United States, Canada, Portugal, the United Kingdom, Slovenia, Australia, Japan, and India (Chalmers et al., 2005a; Feng et al., 2007; Thompson et al., 2007; Trotz-Williams et al., 2006; Xiao et al., 2007b). In areas with less subtype diversity such as Portugal and eastern United States, the IIa15G2R1 subtype is responsible for over 75% of Cryptosporidium infection in calves (Alves et al., 2006b; Xiao et al., 2007b). In Northern Ireland, IIaA18G3R1 was the most common subtype found in cattle, responsible for 55.6% of infections. At least 15 other IIa subtypes occur in calves in Northern Ireland, with IIaA15G2R1 in only 13% of infected calves (Thompson et al., 2007), 23.5 to 27.7% of infected calves in Michigan and Ontario (Peng et al., 2003b; Trotz-Williams et al., 2006), and absent in 18 bovine isolates in Serbia (Misic and Abe, 2007). Some IIa subtypes appear regionally dominant, such as IIaA18G3R1 in Northern Ireland (Thompson et al., 2007), and IIa18G2R1 and IIaA19G2R1 in Florida (Xiao et al., 2007b). Some neighboring areas have been found with a similar distribution of C. parvum subtypes. For example, calves in Michigan and southern Ontario shared five IIa subtypes, three of which (IIaA16G1R1, IIaA16G2R1, and IIaA16G3R1) have not been seen in calves in the lower United States. Frequently, in areas with high C. parvum diversity, one farm has several subtypes of C. parvum circulating in calves (Peng et al., 2003b; Trotz-Williams et al., 2006; Xiao et al., 2007b). The reason for the geographic difference in the distribution of C. parvum subtypes is not clear. However, management practices such as the frequent introduction of new animals can increase the diversity of C. parvum on farms as shown by comparing multilocus subtypes of C. parvum in Turkey and Israel. In Turkey, where animals range over wide areas and interfarm trade of animals is common, C. parvum diversity on each farm is much higher than in Israel, where the traditional kibbutzim farming does not involve much animal trade. Calves in Turkey were more likely to have concurrent infections with mixed C. parvum subtypes (Tanriverdi et al., 2006). The introduction of a new C. parvum subtype can result from transmission of infections among animals housed in close proximity as seen in a zoo setting where the C. parvum IIaA15G2R1 subtype, common in cattle, was found in three Arabian oryxes, three gemsboks, two addaxes, and one eland (Alves et al., 2003b).
V. A.
Molecular Epidemiology of Human Cryptosporidiosis Cryptosporidium Species and Genotypes in Humans
Cryptosporidium parvum was once considered the only Cryptosporidium species to infect humans. Genotyping tools based on DNA sequences of antigen and housekeeping genes identified genotypes 1
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(the human genotype) and 2 (the bovine genotype) within the umbrella of C. parvum, and these eventually became C. hominis and C. parvum, both infectious for immunocompetent and immunocompromised persons (Xiao et al., 2004a; Xiao and Ryan, 2004; Caccio, 2005). The PCR techniques used probably are unable to amplify DNA from some more genetically different Cryptosporidium species. In these earlier studies, a few microscopically positive samples did not generate PCR products when PCR-RFLP techniques based on an undefined genomic sequence, COWP, poly-T, and other genes were used, indicating that unusual Cryptosporidium species might have been present (Bonnin et al., 1996; Widmer et al., 1998; McLauchlin et al., 2000). At the end of the 1990s, SSU rRNA-based genotyping tools revealed the presence of C. canis, C. felis, and C. meleagridis in AIDS patients in the United States, Switzerland, and Kenya, in addition to the more frequently found C. hominis and C. parvum (Morgan et al., 1999a, 2000a; Pieniazek et al., 1999). This observation has been supported by data from France, Portugal, Italy, Thailand, and Peru (Alves et al., 2001; Guyot et al., 2001; Caccio et al., 2002; Gatei et al., 2002b; Tiangtip and Jongwutiwes, 2002). In Peru and Thailand, these three species are responsible for over 20% of Cryptosporidium infections in AIDS patients (Table 5.3). Even immunocompetent persons can be infected with zoonotic species other than C. parvum. Molecular characterization of over 2,000 specimens in the United Kingdom identified 22 cases of C. meleagridis, six cases of C. felis, and one case of C. canis (McLauchlin et al., 2000; Pedraza-Diaz et al., 2000, 2001a, 2001c; Leoni et al., 2006a). Nineteen cases of C. meleagridis infection were identified among 3,100 clinical isolates from England and Wales (Chalmers et al., 2002). Some infected persons were not immunocompromised. HIV-seronegative children in Lima, Peru (Xiao et al., 2001a) and children in Kenya had these Cryptosporidium species (Gatei et al., 2006b). The proportion of infections caused by non-parvum zoonotic Cryptosporidium species, however, was much higher in Peru (about 12%) than in the United Kingdom. At least 16 other cases of C. meleagridis infection have been described in immunocompetent persons in the United Kingdom, France, the Czech Republic, Canada, Japan, and Uganda (Table 5.4). In Peru, where a significant proportion of infections are due to zoonotic Cryptosporidium, there was no significant difference between children and HIV+ adults in the distribution of C. hominis, C. parvum, C. meleagridis, C. felis, and C. canis (Xiao et al., 2001a; Cama et al., 2003). It is likely that other Cryptosporidium species can infect humans under certain circumstances. Cryptosporidium muris-like oocysts were found in two healthy Indonesian girls, but there was no molecular confirmation (Katsumata et al., 2000). A putative C. muris infection was reported in an immunocompromised patient in France based on sequence analysis of a small fragment of the SSU rRNA (Guyot et al., 2001). However, the sequence was more similar to C. andersoni (2-bp differences in a 242-bp region) than to C. muris (8-bp differences in the region). Several confirmed C. muris infections have been documented in AIDS patients in Kenya and Peru, both by PCR-RFLP and sequencing of the SSU rRNA gene (Gatei et al., 2002a, 2006b; Palmer et al., 2003), and a putative human C. muris infection has been seen in India (Muthusamy et al., 2006). More human cases have been associated with the Cryptosporidium cervine genotype, which has been reported in 10 patients in Canada, three patients in the United States, one patient in Slovenia, and one in England (Ong et al., 2002; Blackburn et al., 2006; Feltus et al., 2006; Leoni et al., 2006a; Soba et al., 2006; Trotz-Williams et al., 2006). Other Cryptosporidium species found in humans include C. suis in an HIV+ patient in Lima, Peru, and one patient in England (Xiao et al., 2002a; Leoni et al., 2006a), a C. suis-like parasite in two patients in Canada (Ong et al., 2002), and a C. andersoni-like parasite in three patients in England (Leoni et al., 2006a), and a W17 (chipmunk) genotype in two patients in Wisconsin (Feltus et al., 2006). The C. hominis monkey genotype has been found in two persons in the United Kingdom (Mallon et al., 2003a). Other new Cryptosporidium genotypes will likely be found in humans in future (Chalmers et al., 2002), but these parasites account for a very minor proportion of Cryptosporidium infections in humans. Some unusual Cryptosporidium species may have a broad host range and might emerge as important pathogens in humans when socioeconomic and environmental changes favor transmission. The avian pathogen C. meleagridis is increasingly recognized as an important human pathogen and can experimentally infect a wide range of mammals (Akiyoshi et al., 2003; Huang et al., 2003). In Lima, Peru, and Bangkok, Thailand, C. meleagridis is responsible for 10 to 20% of human cryptosporidiosis cases
b
a
Adults Adults? Adults Adults Adults Adults Adults 4 children, 25 adults Adults Adults Children Children Children Adults Adults Mostly adults Adults Adults Adults Adults
Patient Type
3
14 5 1 1 2 16 34 1 8 1? 38
3
7 1
5 8
17 14 6 56 16 7 8 3 31 204 5 18
34 24 6 76 21 8 10 6 49 302 10 29 1 1? 10 3 3
Gatei et al., 2002b Gatei et al., 2003 Peng et al., 2003a Tumwine et al., 2005 Leav et al., 2002 Meamar et al., 2007 Certad et al., 2006 Navarro-i-Martinez et al., 2006 Ngouanesavanh et al., 2006 Cama et al., 2003 Pieniazek et al., 1999 Xiao et al., 2004a
5 5 1
6 1 3
12 1
1
2
3 6
Reference Alves et al., 2003b Almeida et al., 2006 Morgan et al., 2000a Guyot et al., 2001 Bonnin et al., 1996 Coupe et al., 2005 Muthusamy et al., 2006 Tiangtip and Jongwutiwes, 2002
3
3 2 1 3
16 5 7 22 7 16 9
C. canis
C. felis
C. meleagridis
C. parvum
7 2 2 14 6 25 31 24
C. hominis
29 9 13 46 13 52 48 29
Number of Patients
Including samples from 11 immunocompromised but HIV patients. Including nine HIV-negative children.
Bangkok, Thailand Kenya Malawi Ugandab South Africa Iran Venezuela Colombia Haiti Lima, Peru Atlanta New Orleans
Portugal Portugal Switzerland Francea France France Vellore, India Bangkok, Thailand
Location
Distribution of Common Human-Pathogenic Cryptosporidium Species in HIV-Infected Patients
TABLE 5.3
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563 115 936 6594 71 5 11 25 1 10 20 22 2 83% 16 198 108 4 119 1 13 3 7 5
1263 191 2251 13112 136 39 14 44 11 12 54 29 29
11 178 49 19 5 7 5 0
6 25 44 3 2
26 7 26 17% 6 223 29
64 34 3 18 9
5981
1315
76
662
896
1354
C. parvum
3
2
1 1 2 2?
1
99
22
C. meleagridis
6?
22
6
C. felis
2
1
C. canis
65
21
Mixed Species Reference
Trotz-Williams et al., 2006 Xiao et al., 2004a Feltus et al., 2006 Yagita et al., 2001 Abe et al., 2006 Park et al., 2006 Peng et al., 2001
Mallon et al., 2003a Lowery et al., 2001a Glaeser et al., 2004 Enemark et al., 2002 Hajdusek et al., 2004 Coupe et al., 2005 Ngouanesavanh et al., 2006 Caccio et al., 1999 Soba et al., 2006 Morgan et al., 2000b Chalmers et al., 2005a Learmonth et al., 2004 Ong et al., 2002
Nichols et al., 2006
Smerdon et al., 2003
Hunter et al., 2004a, 2004b
Hunter et al., 2003
McLauchlin et al., 2000; Leoni et al., 2006a Sopwith et al., 2005
136
22 423 150
726
1622
North West England North West England England and Wales England and Wales England and Wales Scotland N. Ireland Switzerland Denmark Czech Republic France France The Netherlands Slovenia Australia Australia New Zealand British Columbia, Canadaa Ontario, Canada United States Wisconsin, USA Japan Japan South Korea China
1005
C. hominis
2414
Number of Isolates
England
Location
Distribution of Common Human-Pathogenic Cryptosporidium Species in Immunocompetent Patients
TABLE 5.4
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a
47 153 35 326 3 3 14 67 2
50 175 37 444 7 62 15 85 4
Includes 9 specimens with the cervine genotype.
India Kenya Malawi Uganda Iran Kuwait Guatemala City, Guatemala Lima, Peru Chile 8 2
0 15 2 85 4 58 1 7
5
1
1
1 2
2
3
2
19
2
Xiao et al., 2001a Neira-Otero et al., 2005
Gatei et al., 2007 Gatei et al., 2006b Peng et al., 2003a Tumwine et al., 2003 Meamar et al., 2007 Sulaiman et al., 2005 Xiao et al., 2004a
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(Xiao et al., 2001a; Gatei et al., 2002b; Cama et al., 2003). Likewise, the increasing number of humans infected with the cervine genotype might be related to its wide range of mammalian hosts.
B.
Role of Animals in Human Cryptosporidiosis Transmission
The host-adapted nature of most Cryptosporidium species indicates that the majority of animal species probably are not important in transmitting cryptosporidiosis to humans. Thus, Cryptosporidium species commonly found in reptiles and most wild mammals have never been detected in humans, and probably have no significant public health importance. However, host adaptation is not strict host specificity. A species or genotype may preferentially infect a species or group of animals, but this does not mean that this parasite cannot infect other animals. Among the over 50 Cryptosporidium species and genotypes described in various animals, only C. parvum, C. meleagridis, C. felis, C. canis, C. muris, C. suis, cervine genotype, and the chipmunk genotype have been found in humans, with C. parvum responsible for most of the infections caused by the zoonotic species (Xiao et al., 2004b; Xiao and Ryan, 2004; Feltus et al., 2006; Leoni et al., 2006a). Many species and genotypes seen in animals and the environment, such as C. bovis, C. baileyi, C. galli, mouse genotype, pig genotype II, and goose genotypes I and II, have never been found in humans. There is no consensus on the human infection potential of the other common bovine species, C. andersoni. SSU rRNA sequences that were more similar to C. andersoni than to C. muris have been found in one patient in France and three patients in England (Guyot et al., 2001; Leoni et al., 2006a). Parasites from different animals and different species from the same animal have vastly different zoonotic potential. Most species of Cryptosporidium infect a limited range of animals, and when the host range or infectivity includes humans, the parasite acquires public health significance. However, the suggestion that the Cryptosporidium mouse genotype is a potential human pathogen because it is closely related to C. parvum (Bajer et al., 2003; Bednarska et al., 2003) requires finding the parasite in human patients, which to date has not occurred. Human-pathogenic Cryptosporidium species and genotypes are scattered all over phylogenetic trees (Xiao et al., 2004b) (Figure 5.2). Thus, the genetic relatedness of Cryptosporidium species is not a good marker for human infectivity. Because C. parvum is the major zoonotic species causing human cryptosporidiosis, and preweaned calves are the major animal host for this parasite, calves are likely the most important source of zoonotic cryptosporidiosis. Contact with infected calves has been implicated as the cause of many cryptosporidiosis outbreaks in veterinary students, research technicians, and children attending agricultural camps and fairs (Preiser et al., 2003; Smith et al., 2004; Kiang et al., 2006). Contamination of food or water by cattle manure has been identified as a cause of several foodborne and waterborne outbreaks of cryptosporidiosis (Millard et al., 1994; Glaberman et al., 2002; Blackburn et al., 2006). Contact with cattle has been implicated as a risk factor in sporadic cases of human cryptosporidiosis in the United States, United Kingdom, Ireland, and Australia in case control studies (Robertson et al., 2002; Goh et al., 2004; Hunter et al., 2004b; Roy et al., 2004). Indeed, massive slaughtering of farm animals and restriction of farm visits during foot-and-mouth disease outbreaks reduced sporadic human C. parvum infections in large communities (Hunter et al., 2003; Smerdon et al., 2003). The contribution of calves in human C. parvum transmission was supported by subtyping sporadic cases in Wisconsin, where most patients were almost exclusively infected with C. parvum (Feltus et al., 2006). Many subtypes found in humans in Wisconsin were previously found in calves in neighboring Michigan and Ontario (Peng et al., 2003b; Trotz-Williams et al., 2006). The role of other domestic and wild ruminants in the transmission of C. parvum to humans is less clear. Cryptosporidium parvum has been found in sheep and some captive ruminants, but the prevalence of C. parvum in these animals is unclear in view of the recent finding of a low prevalence of C. parvum in sheep (Ryan et al., 2005b). Few epidemiologic studies have implicated sheep as a source of human cryptosporidiosis (Duke et al., 1996). However, lambs are sometimes naturally infected with C. parvum, and direct transmission of C. parvum from lambs to children has been confirmed for at least one small outbreak of cryptosporidiosis by subtyping (Chalmers et al., 2005a). One common C. parvum subtype, IIaA15G2R1, has been found in humans, calves, and zoo ruminants in Lisbon, Portugal, suggesting that, under certain circumstances, various domestic and wild ruminants can play a role in transmitting C. parvum to humans (Alves et al., 2003b).
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The role of companion animals in the transmission of human cryptosporidiosis is even less clear. It is been suggested for some time that dogs can be a significant source for human cryptosporidiosis (Bauer, 1994; Enriquez et al., 2001; Robinson and Pugh, 2002). This, however, has been largely based on the observation of direct transmission of C. parvum from calves to humans and the erroneous suggestion that C. parvum is responsible for cryptosporidiosis in all mammals. Only a weak association between cryptosporidiosis in HIV+ persons and contact with dogs was found in the United States (Glaser et al., 1998) or between pediatric cryptosporidiosis and contact with dogs or cats in Guinea-Bissau and Indonesia (Molbak et al., 1994; Katsumata et al., 1998). In England, contact with dogs and cats was not found to be a risk factor for cryptosporidiosis (Goh et al., 2004), and in Australia it was actually a protective factor (Robertson et al., 2002). Dogs and cats are almost exclusively infected with C. canis and C. felis, respectively, whereas humans are mostly infected with C. hominis and C. parvum (Xiao and Ryan, 2004). Thus, the role of companion animals in the transmission of human cryptosporidiosis may be limited. Even though a small number of humans are infected with C. canis and C. felis, the recent finding of a concurrent C. hominis infection in C. canis- or C. felis-infected persons suggest that many of the C. canis and C. felis infections in humans may be due to person-to-person rather than zoonotic transmission (Cama et al., 2006a). A few human-pathogenic Cryptosporidium species have been found in unusual hosts. For example, C. hominis was detected in several calves and sheep in the United Kingdom, United States, Australia, and India (Giles et al., 2001; Ryan et al., 2005b; Smith et al., 2005; Feng et al., 2007), C. suis has been found in a calf in the United States (Fayer et al., 2006) and a few lambs in Australia (Ryan et al., 2005b), C. meleagridis was seen in one dog in the Czech Republic (Hajdusek et al., 2004), and the C. canis dog genotype was seen in a fox (Zhou et al., 2004a). The role of these animals in the transmission of these species to humans is probably minimal. Mechanical carriage of C. hominis and C. parvum oocysts has been reported in a few Canada geese (Graczyk et al., 1998; Zhou et al., 2004b). However, Canada geese are normally infected with two unique Cryptosporidium genotypes, goose genotypes I and II (Zhou et al., 2004b).
C.
Molecular Epidemiology of Endemic Human Cryptosporidiosis
Molecular characterization of Cryptosporidium isolates reveals the complexity of cryptosporidiosis epidemiology. Among the five common Cryptosporidium species in humans, C. parvum and C. hominis are responsible for greater than 90% of human cases of cryptosporidiosis in most areas (Xiao and Ryan, 2004). Geographic differences exist in the disease burdens attributable to these two species. In the United Kingdom, other parts of the Europe, and New Zealand, C. parvum is responsible for slightly more infections than C. hominis (Homan et al., 1999; McLauchlin et al., 2000; Guyot et al., 2001; Chalmers et al., 2002; Alves et al., 2003b; Fretz et al., 2003; Hajdusek et al., 2004; Learmonth et al., 2004; Leoni et al., 2006a). In contrast, C. hominis is responsible for far more infections than C. parvum in the United States, Australia, Japan, and developing countries where genotyping studies have been conducted (Peng et al., 1997, 2003a; Morgan et al., 1998; Sulaiman et al., 1998; Ong et al., 1999, 2002; Xiao et al., 2001a; Leav et al., 2002; Tiangtip and Jongwutiwes, 2002; Cama et al., 2003; Gatei et al., 2003, 2006b; Tumwine et al., 2003, 2005; Chalmers et al., 2005a; Bushen et al., 2006; Das et al., 2006; Muthusamy et al., 2006). Interestingly, in the highly urbanized Kuwait City, almost all cryptosporidiosis cases in children are caused by C. parvum (Sulaiman et al., 2005), which may also be the case in Saudi Arabia. This geographic difference in the distribution of C. parvum and C. hominis in humans is true in both immunocompetent (Table 5.4) and immunocompromised individuals (Table 5.3). Major differences in the transmission routes may be responsible for the differences in Cryptosporidium species distribution. This is supported by results of studies in the United Kingdom, which reported that C. hominis infection was more common in patients with a history of foreign travel (McLauchlin et al., 2000; Goh et al., 2004; Hunter et al., 2004b). Not surprisingly, geographic differences in the distribution of Cryptosporidium genotypes can occur within a country (McLauchlin et al., 1999, 2000; Learmonth et al., 2004). Thus, C. hominis infection is generally more common in urban areas, and C. parvum more common in rural areas. It seems likely, but remains unproven, that the high prevalence of C. parvum in humans in some regions of the world may be due, in part, to the intensive husbandry practiced for
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ruminants and the associated high concentrations of young animals at these feeding operations. In the United States, even though C. hominis is usually more common than C. parvum in humans (Xiao et al., 2004a), most human cryptosporidiosis cases in the dairy state Wisconsin are attributable to C. parvum (Feltus et al., 2006). Indeed, restriction of farm visits and culling of farm animals during a foot-andmouth disease outbreak in England have greatly reduced the occurrence of cryptosporidiosis due to C. parvum (Hunter et al., 2003; Smerdon et al., 2003). It is possible that this and other factors may have permanently changed the transmission of cryptosporidiosis in North West England (Sopwith et al., 2005). In recent years, C. hominis has been more prevalent than C. parvum in humans in England and Wales (Nichols et al., 2006) Seasonal differences in the distribution of C. parvum and C. hominis have been reported. In the United Kingdom and New Zealand; the spring increase in the reported cryptosporidiosis cases was mostly due to C. parvum, whereas the autumn increase was largely due to C. hominis (McLauchlin et al., 2000; Learmonth et al., 2003, 2004; Hunter et al., 2004b), suggesting that seasonal differences may exist in the relative importance of specific transmission routes. It has been speculated that the increase in C. parvum in spring is due to lambing, calving, and runoff from spring rains, and the autumn C. hominis peak in these countries is likely the result of increased recreational water activities and international travel during late summer and early autumn (Goh et al., 2004; Hunter et al., 2004b). The transmission of human cryptosporidiosis has been examined in several areas using GP60-based subtyping tools. Results of these studies have revealed the complexity of Cryptosporidium transmission in endemic areas. This is evident by the existence of many C. hominis and C. parvum subtype families in each endemic area. Thus, three to four C. hominis subtype families were seen in humans in India, Peru, New Orleans, Malawi, South Africa, Kuwait, and Portugal, even though only one or two C. parvum subtype families were usually seen (Leav et al., 2002; Alves et al., 2003b, 2006b; Peng et al., 2003a; Xiao et al., 2004a, 2004b; Xiao and Ryan, 2004; Sulaiman et al., 2005; Gatei et al., 2007). In developing countries, the complexity of transmission is frequently reflected by the existence of many subtypes within C. hominis subtype families Ia and Id (Leav et al., 2002; Peng et al., 2003a; Gatei et al., 2007). The high C. hominis heterogeneity in developing countries is likely an indicator of stable cryptosporidiosis transmission in the area. In contrast, the heterogeneity in C. hominis in industrialized nations is generally smaller, reflected by the common occurrence of subtype family Ib and/or the low heterogeneity in subtype families Ia and Id (Glaberman et al., 2002; Alves et al., 2003b, 2006b; Xiao et al., 2004b; Chalmers et al., 2005a). The predominance of two major subtypes (IIaA15G2R1 and IIdA20G1) in children in Kuwait City is likely the result of cryptosporidiosis becoming recently endemic with unstable cryptosporidiosis transmission in the area (Sulaiman et al., 2005). Two major GP60 subtype families, IIa and IIc, are responsible for most C. parvum infections in humans. The distribution of the two subtype families, however, varies by geographic regions. In the rural areas in the United States and in Europe, many C. parvum isolates belong to subtype family IIa, which is the major zoonotic Cryptosporidium subtype (Glaberman et al., 2002; Alves et al., 2003b, 2006b; Chalmers et al., 2005a; Feltus et al., 2006). In urban areas in the United States and in developing countries, however, IIa subtypes are rarely seen in humans. Instead, the anthroponotic IIc subtype family is responsible for most human C. parvum infections in these areas (Leav et al., 2002; Peng et al., 2003a; Xiao et al., 2004b; Xiao and Ryan, 2004; Akiyoshi et al., 2006). In European countries such as Portugal and the United Kingdom, both IIa and IIc are fairly common in humans (Alves et al., 2003b, 2006b). In some regions such as Lima, Peru, the IIc subtype family is the only C. parvum in humans, whereas in other developing countries such as Malawi and Kenya, IIe (another anthroponotic C. parvum subtype family) is seen in humans in addition to IIc (Peng et al., 2003a; Xiao et al., 2004a). In Uganda, even though IIc subtypes are the dominant C. parvum in children, several new subtype families are seen (Akiyoshi et al., 2006). An unusual zoonotic C. parvum subtype family IId is seen in some humans in Portugal (Alves et al., 2003b, 2006b). All these are likely indicators of differences in the role of zoonotic parasites in the transmission of C. parvum among geographic areas. However, anthroponotic transmission of seemingly zoonotic C. parvum subtype families can occur. For example, nearly all cryptosporidiosis cases in children in Kuwait City are attributable to two zoonotic C. parvum subtype families, IIa and IId, even though zoonotic transmission is likely not important in the highly urbanized area (Sulaiman et al., 2005).
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The four common C. hominis subtype families Ia, Ib, Id, and Ie have been found in humans in many areas (Xiao et al., 2004a; Xiao and Ryan, 2004). Nevertheless, there are geographic differences in the distribution of these subtype families. For example, Ib was a predominant C. hominis subtype family in humans in C. parvum-endemic areas such as Portugal, United Kingdom, and Australia (Glaberman et al., 2002; Alves et al., 2003b, 2006b; Chalmers et al., 2005a). Both Ib and Id were common in developing countries such as Malawi, South Africa, India and Peru (Leav et al., 2002; Peng et al., 2003a; Xiao et al., 2004a; Gatei et al., 2007). In New Orleans, Ib and Ia were both common, and Id subtypes were absent (Xiao et al., 2004b). Children in South Africa were commonly infected with If (erroneously named as Ie in the publication), a subtype family not seen in most other studies (Leav et al., 2002) but was reported in HIV+ adults in India (Muthusamy et al., 2006). The latter, however, was based on results of RFLP analysis rather than DNA sequencing. Because there is some confusion in naming subtype families Ie and If, it is not clear whether the If subtype family reported by Muthusamy et al. (2006) was actually Ie. Likewise, subtype family Ie was not seen in cryptosporidiosis in South African children (Leav et al., 2002). Within each subtype family, one subtype is frequently seen in certain areas but not in others. For example, there are only two common subtypes within the C. hominis subtype family Ib: IbA9G3 and IbA10G2. The former is commonly seen in Malawi, Kenya, India, and Australia, whereas the latter is commonly seen in South Africa, Peru, the United Kingdom, and the United States (Glaberman et al., 2002; Leav et al., 2002; Alves et al., 2003b, 2006b; Xiao et al., 2004b; Chalmers et al., 2005a; Gatei et al., 2006a, 2007). Frequently, only one of the two subtypes is seen in humans in one area. Likewise, within the C. hominis subtype family Ie, humans in most areas are infected with IeA11G3T3¸ with the exception of New Orleans, USA, and Adelaide, Australia, where IeA12G3T3 is seen (Xiao et al., 2004b; Chalmers et al., 2005a). Genotyping has been used in the study of the development of immunity against Cryptosporidium infection. Although it was suggested that immunocompromised persons are more susceptible to infections with zoonotic Cryptosporidium species (Gibbons et al., 2001; Guyot et al., 2001; Tumwine et al., 2005), results of genotyping in Lima, Peru, indicate that this may not be necessarily the case. In that, C. meleagridis, C. felis, and C. canis were found in both AIDS patients and children, responsible for 12.5% of infection in children and 17.4% of infections in AIDS patients (Xiao et al., 2001a; Cama et al., 2003). Molecular analysis of longitudinal samples from 16 Peruvian children with multiple cryptosporidiosis episodes indicated that immunity against both homologous and heterologous Cryptosporidium species and genotypes was short lived, with time intervals between infections of about one year. As might be expected, sequential infections with heterologous Cryptosporidium species were more common than sequential infections with homologous Cryptosporidium species; ten events of heterologous infections versus six events of homologous infections with C. hominis were seen in these children, even though C. hominis was the most common Cryptosporidium species in the community, responsible for 70% of all infections (Xiao et al., 2004a). Subtyping was done on samples from sequential infections for three of the six children with multiple episodes of infections with homologous genotypes (C. hominis). One child (P5076) had one episode of infection due to subtype family Id and two subsequent episodes due to Ie, one child (E392) had the first episode of infection due to Ib and the second one due to Ie, and one child (K283) had one infection due to Id and one due to Ib. These results suggest that multiple episodes of infections with homologous genotypes can be in fact due to heterogeneous subtypes of the parasite (Xiao et al., 2004a).
D.
Molecular Epidemiology of Epidemic Human Cryptosporidiosis
Although many Cryptosporidium species and genotypes are found in humans, to date only C. parvum and C. hominis have been linked to outbreaks of cryptosporidiosis (Peng et al., 1997; Ong et al., 1999; McLauchlin et al., 2000; Xiao et al., 2004a; Chalmers et al., 2005a; Smith et al., 2006a). In the United States, between 1993 and 2006, C. hominis and C. parvum were identified as the major cause in 20 and 7 of 28 food- and waterborne outbreaks, respectively, and in one outbreak, both Cryptosporidium species were detected (Table 5.5). However, only a small number of samples were analyzed in many of the outbreaks, especially the earlier ones. In 13 waterborne outbreaks in England during 1994–1999, where
1993
1993 1994 1995 1995 1997 1997 1998 1998 2000 2000 2000 2000 2000 2000 2001 2001 2001
2001
2002
Maine
Las Vegas, NV Gainesville, FL
Cobb Co., GA Minnesota Spokane, WA Austin, Texas Washington, DC Charleston, SC Canon, CO Omaha, NE
Powell, OH
N. Ireland N. Ireland
N. Ireland
E. Peoria, IL N. Battleford, Saskatchewan
South Burgundy, France
San Antonio, TX
Year
Milwaukee, WI
Outbreak
Waterborne
Waterborne
Waterborne Waterborne
Waterborne
Waterborne Waterborne
Waterborne
Swimming in a swimming pool
Drinking water
Drinking water Drinking water contaminated by ingress of human sewage from a septic tank Drinking water contaminated by wastewater from a blocked sewer drain Attendance to a water park Drinking water: intake downstream wastewater discharge
Swimming in a swimming pool
Drinking water: slaughterhouse discharge, pasture runoff, human sewage? Apple cider from manure-contaminated apples Lake/drinking water Day camp: fecal contamination of tap water? Attendance to a water park Water fountain in zoo: diapered children? Holiday party: green onion? Drinking water: sewage seepage into well University cafeteria, food handler Swimming in a swimming pool Swimming in a swimming pool Swimming in a swimming pool
Contamination source
1
23
11 10
44
33 32
31
3 5 7 5 25 6 3 24
4 6
1
5
IIaA15G2R1
0/1
1/0
23/0
11/0 17/0
36/8
0/33 32/0
31/31a
3/0 0/5 7/0 4/1 25/0 6/0 0/3 24/0
IbA10G2 (36/44), IIaA18G3R1 (8/44) IaA14R3 (11/11) IdA19 (7/13), IbA10G2 (4/13), mixed (2/13) IbA10G2 (21/23), IdA24 (1/23), IIaA15G2R1 (1/23) IbA10G2 (1/1)
IdA17G1 (20/21), IIaA15G2R1 (1/21) IIaA18G3R1 (30/30) IbA10G2 (31/31)
IbA10G2 (15/15) IaA24R4 (6/6) IIcA5G3 (3/3) IaA24R4 (12/12)
IbA10G2 (3/3) IIaA15G2R1 (4/4) IdA18 (7/7)
IbA10G2 (3/3) IbA9G3 (3/3)
IbA10G2 (4/4)
5/0
3/1 6/0
Subtype in Patient Sample
Species C. hominis/ C. parvum
IbA10G2 in implicated blocked
IaA24R4 (1) in swimming pool
Subtype in Environmental Sample
142
Waterborne Waterborne Foodborne Waterborne Foodborne Waterborne Waterborne Waterborne
Waterborne Waterborne
Foodborne
Waterborne
Route
Number of Patients
Distribution of C. parvum and C. hominis GP60 Subtypes in Foodborne and Waterborne Outbreaks of Human Cryptosporidiosis in North America and Europe
TABLE 5.5
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2005 2005 2006 2006
2006
2006 2006
Seneca Lake, NY
Oregon Nassau, FL
Anderson, SC
Tazewell Co., IL
Lone Tree, CO Ashtabula Co., OH Waterborne Foodborne
Waterborne
Waterborne
Waterborne Unknown
Waterborne
Waterborne Waterborne Waterborne Waterborne Waterborne
Waterborne Foodborne
Swimming in a swimming pool Drinking apple cider
Summer camp, swimming in camp pool or visiting a water park
Swimming in a swimming pool Travel to Ireland, waterborne or foodborne Swimming in a swimming pool
Attendance to a splash park
Swimming in a swimming pool in Spain Attendance to a water park Swimming in a swimming pool Swimming in a swimming pool Swimming in a swimming pool
Indoor water park Drinking ozonated apple cider
Mixed infection of both species in all patients examined.
2003 2004 2004 2004 2005
Ireland San Luis Obispo Co., CA Colorado Auglaize Co., OH Hamilton County, OH
a
2003 2003
Douglas Co., KS Stark Co., OH
5 2
4
4
4 5
9 6 1 17 7
49 12
IbA10G2 (5/5) IdA14 (2/2)
IbA10G2 (4/4)
4/0
5/0 2/0
IaA28R4 (4/4)
IIaA16G1R1b (5/5)
IIcA5G3h (9/9) IIcA5G3 (6/6) Negative IdA15G1 (17/17) IaA28R4 (6/7), IdA15G2 (1/7)
IdA17 (48/48) IIaA15G2R1 (4/8), IIaA17G2R1 (4/8)
4/0
0/4 0/5
0/9 0/6 1/0 17/0 7/0
49/0 0/11
IaA28R4 in splash zoon; IIaA18G3 in lap pool Negative for camp pool, IIaA15G2R1 in another
IbA10G2 (2)in water in tanks 1 and 2
IaA28R4
IIaA17G2R1 in implicated apple cider
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larger numbers of samples were analyzed, C. hominis was the predominant cause for 7 outbreaks, C. parvum for 4 outbreaks, and both species for 2 outbreaks (McLauchlin et al., 2000). Similarly, in 11 waterborne outbreaks of cryptosporidiosis in England and Wales during 2000–2003, C. hominis was the major cause in 6 outbreaks and C. parvum in 4 outbreaks, with both species as the cause in one outbreak (Smith et al., 2006a). Thus, even in C. parvum-endemic areas, C. hominis is responsible for more waterborne outbreaks of cryptosporidiosis than C. parvum. Subtyping tools for C. hominis and C. parvum have been used successfully in cryptosporidiosis outbreak investigations. The ds-RNA subtyping tool was used in the investigation of one waterborne and two foodborne outbreaks in the United States. Isolates from each outbreak analyzed had identical ds-RNA sequences, and three outbreaks were caused by three different subtypes (Xiao et al., 2001b). In contrast, in 43 specimens from sporadic infections, a total of 15 subtypes were found, none of which were identified in the outbreak cases (Xiao et al., 2001b). This is very different from results of outbreak investigations in England, where one single C. hominis subtype was seen in all three waterborne outbreaks, and eight intrafamilial outbreaks investigated, and another C. parvum subtype was the major cause of two waterborne outbreaks and seven intrafamilial outbreaks, even though multiple subtypes of C. hominis and C. parvum were seen in sporadic cases (Leoni et al., 2003b). The role of subtyping is best demonstrated by the GP60-based subtyping analysis. As noted earlier, this tool divides C. hominis and C. parvum into nine major subtype families (Figure 5.1), each of which contains multiple subtypes. Thus far, three C. hominis (Ia, Ib, and Id) and two C. parvum (IIa and IIc) subtype families have been identified in cryptosporidiosis outbreaks in the United States, Canada, United Kingdom, and France (Glaberman et al., 2002; Xiao et al., 2004a; Chalmers et al., 2005a; Cohen et al., 2006). The most common subtype family is Ib, especially the subtype IbA10G2, which has been found in nearly half (9/19) of the C. hominis outbreaks investigated in the United States. IbA10G2 has been responsible for waterborne cryptosporidiosis outbreaks in the United Kingdom, Canada, and France (Glaberman et al., 2002; Ong et al., 2005; Cohen et al., 2006). For most of the outbreaks examined, all isolates from the same outbreak had the same GP60 subtype (Table 5.5). Genotyping and subtyping are useful in the detection and differentiation of outbreaks. During the investigation of the apple-cider-associated outbreaks of cryptosporidiosis in 2003 in Ohio, all patient stool specimens collected early in the outbreak had C. parvum, with either subtype IIaA15G2R1 or IIaA17G2R1 (Blackburn et al., 2006). In contrast, among the specimens collected later, five had C. hominis, one had both C. hominis and C. parvum, and one had the Cryptosporidium cervine genotype. All the C. hominis specimens were from children attending a daycare center in the outbreak area, and all belonged to the subtype IbA9G3. Interestingly, the patient with mixed C. hominis and C. parvum infection had all three subtypes, IbA9G3, IIaA15G2R1, and IIaA17G2R1. Thus, two outbreaks of cryptosporidiosis occurred roughly during the same period with very different causes, and at least one patient was involved in both outbreaks. Likewise, both C. hominis and C. parvum were seen among submitted specimens during the investigation of the drinking-water-associated cryptosporidiosis outbreak in Northern Ireland in 2001 (Glaberman et al., 2002). All specimens from 36 patients living in the outbreak area had C. hominis subtype IbA10G2, whereas all eight specimens from areas not affected by the outbreak had C. parvum subtype IIaA18G3R1, which is the most commonly identified Cryptosporidium subtype in Northern Ireland (Thompson et al., 2007). Interestingly, the parasites involved in two of the swimming-pool-associated cryptosporidiosis outbreaks in summer of 2000, in Nebraska and South Carolina, were all of the same C. hominis subtype (IaA24R4), although this subtype appears to be rare (Table 5.5). There were no epidemiologic data available to determine whether the two outbreaks were linked. Subtyping is very useful in tracking the source of infection or contamination sources during waterborne outbreak investigations. 1. Cryptosporidium hominis subtype (IaA24R4) oocysts found in three sand samples from the filter bed of the swimming pool involved in a Nebraska outbreak in 2000 matched the subtype in stool specimens from the 10 outbreak patients analyzed.
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2. Similarly, in an Ohio swimming pool outbreak in 2000, the C. hominis subtype IdA17G1 found in four sand samples from the filter bed was identical to the human subtype found in specimens from all outbreak patients. 3. In a drinking-water-associated cryptosporidiosis outbreak in Northern Ireland in 2001, a water sample from the implicated blocked wastewater drain had the same subtype as found in specimens from outbreak patients. 4. Cryptosporidium hominis subtype IaA28R4 was found in both the swimming pool and patient specimens from two swimming-pool-associated outbreaks of cryptosporidiosis in Ohio in 2005 and in South Carolina in 2006. 5. In a 2006 Illinois waterborne outbreak of cryptosporidiosis at a summer camp, children potentially got exposed by swimming in the camp pool or visiting a water park. All outbreak cases had C. hominis IbA10G2, whereas the backwash sample from the water park pool was strongly positive for C. parvum subtype IIaA15G2R1, which exonerated the water park as the site of exposure (Table 5.5). G60-based subtyping has been used in identifying the source of contamination in foodborne outbreaks of cryptosporidiosis. In a 1998 Washington D.C. foodborne cryptosporidiosis outbreak, three specimens from the epidemiologically implicated food handler had the same C. hominis subtype (IbA10G2) as did outbreak patients (Quiroz et al., 2000). In the 2003 apple-cider-associated cryptosporidiosis outbreak in Ohio, one of the two C. parvum subtypes detected in patient specimens, IIaA17G2R1, was found in the leftovers of the implicated apple cider (Blackburn et al., 2006). This subtype is reasonably rare in the eastern United States, having only been reported in calves commonly in Ohio and occasionally in Vermont (Xiao et al., 2007b). Thus, cattle were likely the source of contamination of apples with Cryptosporidium oocysts, which was already implicated as the cause (IIaA15G2R1) of an earlier apple-cider-associated cryptosporidiosis outbreak in Maine (Millard et al., 1994) (Table 5.5).
E.
Biological, Clinical, and Epidemiologic Differences among Cryptosporidium Species
The clinical and epidemiologic significance of various Cryptosporidium species and genotypes in humans is not yet clear. Biologically, C. parvum and C. hominis differ from each other in host specificity. Cryptosporidium parvum infects humans and ruminants in natural situations and mice in cross-transmission experiments, whereas C. hominis does not readily infect laboratory animals or calves (Peng et al., 1997). Nevertheless, the number of isolates tested is small, and natural C. hominis infections have been reported in a few calves, lambs, kids and a dugong (Morgan et al., 2000c; Giles et al., 2001; Tanriverdi et al., 2003; Smith et al., 2005; Park et al., 2006; Feng et al., 2007). The only well-established laboratory animal model for both C. parvum and C. hominis is the gnotobiotic pig model, which uses very low infection doses and allows direct comparison of these two species within the same host (Widmer et al., 2000; Pereira et al., 2002; Chapter 19). In gnotobiotic pigs, C. parvum and C. hominis differ from each other in the prepatent period, infection site, and disease severity. Isolates of C. hominis have longer prepatent periods than C. parvum isolates (8.8 versus 5.4 days). With the few isolates tested, C. parvum infects the entire small and large intestine, whereas C. hominis infects mostly ileum and colon, which have higher number of parasites per villous when infected with C. hominis. Cryptosporidium parvum–infected pigs develop moderate-to-severe disease, whereas C. hominis-infected pigs have only mild-to-moderate disease. Moderate-to-severe villous/mucosal attenuation with lymphoid hyperplasia is usually seen throughout the intestine of C. parvum-infected pigs. Lesions in C. hominis-infected pigs are mild to moderate and restricted to the ileum and colon (Pereira et al., 2002). An immunosuppressed Mongolian gerbil model for both C. parvum and C. hominis has also been described, which requires the use of higher infection doses than the gnotobiotic model. No difference in oocyst shedding was observed between the two Cryptosporidium species in this model (Baishanbo et al., 2005).
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The significance of these findings in gnotobiotic pigs is not entirely clear. Because bovine Cryptosporidium isolates infect the entire small and large intestines in experimentally infected calves, lambs, pigs, and mice, the biologic differences between C. hominis and C. parvum in gnotobiotic pigs may represent intrinsic differences between these two genetically different parasites. These two patterns of infections in gnotobiotic appear to parallel the ileocolonic infection versus panenteric infection seen in AIDS patients infected with Cryptosporidium species. In the latter, panenteric infection is associated with more severe villous atrophy, lamina propria inflammatory infiltration, and clinical symptoms, and poorer survival than ileocolonic infection. It is not clear at this moment whether these two patterns of cryptosporidiosis in AIDS patients are due to infections with two different species. Results of recent genotyping studies nevertheless support the theory that C. hominis and C. parvum behave differently in humans. Of 194 sporadic cases of cryptosporidiosis in the United Kingdom, 52.7% (39/74) of samples with C. hominis were scored microscopically for 2+ or 3+, but only 36.7% (44/120) of samples with the C. parvum were scored for 2+ or 3+, indicating that humans infected with C. hominis may shed more oocysts than those infected with C. parvum (McLauchlin et al., 1999). Similarly, in children in Peru, stools with C. hominis had a significantly higher mean oocyst score than those with the zoonotic genotypes (1.7 versus 1.3 out of 3, p = 0.02). Infections with C. hominis had a significantly longer duration of oocyst shedding than those with the zoonotic genotypes in the longitudinally followed cohort of Peruvian children (13.9 versus 6.4 days, p = 0.004) (Xiao et al., 2001a). Likewise, oocystshedding intensity was higher in Brazilian children infected with C. hominis than C. parvum (Bushen et al., 2007). In addition to the differences in host specificity and oocyst shedding, C. parvum and C. hominis differ from each other in pathogenicity and clinical presentations. In sporadic cryptosporidiosis in England, C. hominis, but not C. parvum, was associated with an increased risk of nonintestinal sequelae such as joint pain, eye pains, recurrent headache, dizzy spells, and fatigue (Hunter et al., 2004a). In AIDS patients in Lima, Peru, only infections with C. canis, C. felis, or subtype family Id of C. hominis were significantly associated with diarrhea, and infections with C. parvum were associated with chronic diarrhea and vomiting. In contrast, infections with C. meleagridis and Ia and Ie subtype families of C. hominis were usually asymptomatic. These results demonstrate that different Cryptosporidium genotypes and subtype families are linked to different clinical manifestations (Cama et al., 2007). Because of the differences in pathogenicity, C. hominis and C. parvum seemingly have different nutritional effects on infected children. In Brazil, height-for-age (HAZ) Z-scores showed significant declines within 3 months of infection for children infected with either C. hominis or C. parvum. However, in the 3-6 month period following infection, only C. hominis-infected children continued to demonstrate declining HAZ score, and those with asymptomatic infection showed even greater decline (p = 0.01). Thus, C. hominis is associated with greater growth shortfalls even in the absence of symptoms (Bushen et al., 2007).
VI. Cryptosporidium Contamination Source Tracking Several genotyping tools have been used to determine whether Cryptosporidium oocysts found in water are from human-infective species and what the likely contamination source is (Table 5.6). In one of the earlier studies, an SSU rRNA-based nested PCR-RFLP method was used in the detection and differentiation of Cryptosporidium oocysts present in storm water (Xiao et al., 2000). Twelve wildlife genotypes of Cryptosporidium were detected in 27 positive samples of storm water collected from a stream that contributes to the New York City Water Supply system. Only the W4 (cervine) genotype has ever been found in humans and, therefore, most Cryptosporidium oocysts found in storm runoff probably do not infect humans. Twelve of the 27 PCR positive samples had multiple genotypes (Xiao et al., 2000). More recently, the molecular characterization has been extended to two other streams in the watershed. A total of 22 Cryptosporidium genotypes were found in storm water collected from the three streams, almost all of which represent wildlife genotypes, and only two common genotypes (W4 and W17, or the cervine and chipmunk genotypes) have been found in a few humans. The distribution of Cryptosporidium genotypes in storm water varied between streams and season (Jiang et al., 2005b). In both the initial
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and expanded studies, the sensitivity of the SSU rRNA–based nested PCR was much higher than microscopy (Xiao et al., 2000, 2006b). The same technique used in the analysis of surface water samples collected from Maryland, Wisconsin, Illinois, Texas, Missouri, Kansas, Michigan, Virginia, Iowa, and Washington in the United States produced quite different results (Xiao et al., 2001c). Of 55 samples, 25 were PCR positive. Only C. parvum, C. hominis, C. andersoni, and C. baileyi were found, two of which are known human pathogens, and humans and farm animals were major contributors of contamination (Xiao et al., 2001c). There was very good agreement between genotyping results and environmental settings. In one watershed, C. parvum was found downstream of farms, whereas both C. parvum and C. hominis were found downstream of urban wastewater discharge sites. Similar results in genotype distribution were obtained from another 113 surface-water samples in the United States by the same researchers (Xiao et al., 2006a), and by others in surface-water samples in Canada and Portugal (Ruecker et al., 2005; Alves et al., 2006a) (Table 5.6). Other SSU rRNA–based PCR-sequencing tools have been successfully used in the differentiation of Cryptosporidium oocysts in surface-water samples (Jellison et al., 2002; Ward et al., 2002; Coupe et al., 2006; Hashimoto et al., 2006; Hirata and Hashimoto, 2006; Masago et al., 2006). Sequences of C. andersoni and, presumably, C. parvum and C. baileyi, were obtained from seven samples of surface water from a watershed in Massachusetts (Jellison et al., 2002). Analysis of 17 positive surface-water samples from Germany and Switzerland found seven Cryptosporidium genotypes, with C. parvum, C. muris, and C. andersoni as the most prevalent, and three samples having three new wildlife genotypes and C. baileyi (Ward et al., 2002). Similar results were obtained with SSU rRNA-based nested PCR, seminested PCR, or real-time PCR in the United Kingdom, France, and Japan (Coupe et al., 2006; Hashimoto et al., 2006; Hirata and Hashimoto, 2006; Masago et al., 2006; Nichols et al., 2006). These and earlier studies showed that humans, cattle, and wild animals are major contributors of oocyst contamination of surface water; the relative contribution of each animals depends on the environmental settings; and many of the oocysts found in source water are not from human-pathogenic species and genotypes (Table 5.6). An SSU-based nested-PCR tool was used to genotype Cryptosporidium oocysts in 14 finished water samples in Scotland; only C. hominis was found (Nichols et al., 2003). SSU rRNA-based genotyping tools have been used in analysis of wastewater samples. Cryptosporidium parvum, C. hominis, C. andersoni, C. muris, C. canis, C. felis, and the cervine genotype were found in 12 of 49 raw urban wastewater collected from a treatment plant in Milwaukee, Wisconsin, with C. andersoni the most common species (Xiao et al., 2001c). These species were found in two subsequent studies conducted in the same wastewater treatment plant, with C. andersoni and C. hominis most prevalent (Zhou et al., 2003; Xiao et al., 2006a). Similar results were obtained in wastewater samples characterized in Germany, Switzerland, and Japan (Ward et al., 2002; Hashimoto et al., 2006; Hirata and Hashimoto, 2006), except for the absence of C. andersoni. In Milwaukee, subtyping done nearly a decade after the massive waterborne cryptosporidiosis outbreak in 1993 found that the original C. hominis subtype involved in the outbreak, IbA10G2, was still the major C. hominis subtype in the community (Zhou et al., 2003). In an evaluation study, the sensitivity of two SSU rRNA-based nested PCRs in the detection of Cryptosporidium oocysts in water was higher than that of two COWP- and DHFR-based PCR tools (Nichols et al., 2006). Between the two SSU rRNA-based nested PCR assays tested, the newly developed technique was more sensitive than the commonly used technique. However, the sequence of the reverse primer used in primary PCR for the latter technique was from an earlier publication (Xiao et al., 1999c), which had three nucleotide errors, corrected in the subsequent publications (Xiao et al., 2000, 2001c). In other studies, genotyping techniques based on other antigen and housekeeping genes such as TRAPC2 and HSP-70 have been used successfully in characterizing Cryptosporidium oocysts in surface water samples (Di Giovanni et al., 1999; Karasudani et al., 2001; Lowery et al., 2001b, 2001c; LeChevallier et al., 2003; Hanninen et al., 2005). The diversity of Cryptosporidium species found, however, were generally smaller than that seen in studies conducted with SSU rRNA-based techniques, with only C. parvum and/or C. hominis detected in most of the studies (Table 5.6). Although oocysts of all Cryptosporidium species can potentially appear in water, only a few are known human pathogens. Even among the latter, not all have the same infectivity for humans. Very limited
SSU rRNA
TRAP-C2 SSU rRNA and TRAP-C2 SSU rRNA
HSP70 SSU rRNA SSU rRNA
SSU rRNA HSP70 SSU rRNA
IMS-nested PCRa
IMS-PCRa
IMS-nested PCR-RFLP
RT-PCRa
IMS-nested PCR IMS-nested PCR
IMS-nested PCR IMS-CC-PCRa
IMS-nested PCR
HSP70
IMS-nested PCR-RFLP
CC-PCR
a
Method
Gene Target
0.05
Unknown 10
40–80 2 or 20
2.5–10
Finished water: 14/14 Source water: 22/560 Filter backwash: 9/121 Raw wastewater: 50/179
Surface water: 7/78 Surface water: 24/60; Wastewater: 6/8
River water: 2/6
Surface water: 25/55 Raw wastewater: 12/49
Surface and finished water: 11/214 River water and sewage effluent: 2/10
Source water: 6/122 Backwash: 9/121 Storm water: 27/29
Number of Positive/Number of Samples
Surface water: C. andersoni (5), C. parvum (10), C. hominis and C. parvum (9), C. hominis and C. baileyi (1) Wastewater: C. andersoni (5), C. canis (1), C. muris (1), C. felis (1), cervine genotype (1), C. andersoni and C. hominis (1), C. andersoni and C. parvum (1), C. andersoni and C. muris (1) C. parvum (1), and C. parvum and C. meleagridis (1) C. parvum? (3), C. andersoni (3), C. baileyi? (1) Surface water: C. parvum (5), C. muris (5), C. andersoni (3), C. baileyi (1), 3 new genotypes (3), dinoflagellates (7) Wastewater: C. hominis (3), C. parvum (2), C. muris (1) C. hominis (12), unknown (2) Source water: C. parvum (19), C. hominis (2), new genotype (1) C. hominis (24), C. andersoni (23), C. parvum (5), C. muris (4), mouse genotype (1), cervine genotype (6)
C. parvum
C. parvum
7 sequence types, probably from C. parvum, C. hominis, and the mouse genotype 12 species/genotypes, all from wildlife
Species or Genotype
Nichols et al., 2003 LeChevallier et al., 2003 Zhou et al., 2003
Jellison et al., 2002 Ward et al., 2002
Karasudani et al., 2001
Xiao et al., 2001c
Lowery et al., 2001c
Lowery et al., 2001b
Xiao et al., 2000
Di Giovanni et al., 1999
Reference
148
Surface water: 10–63.1 Wastewater: 0.01–0.05
500
189–224; IMS done on Percoll-sucrose concentrates 500–1000
10
Volume of Water (L)
Genotyping Cryptosporidium in Natural Water by PCR-Based Techniques
TABLE 5.6
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SSU rRNA
SSU rRNA
SSU rRNA
SSU rRNA SSU rRNA SSU rRNA
IMS-nested PCR
IMS-real-time PCR
IMS-semi-nested PCR
IMS-nested PCR
IMS-nested PCR
IMS-nested PCR Surface water: 10 Raw wastewater: 0.05
Unknown
20
Single oocyst from 15 wastewater and 3 river water samples
23.8–66.1
1–10
Surface water: 10 Wastewater and sludge: 0.5–2 50
Microscopy-positive surface water: 6/15 Surface, ground and finished water: 40/123 Surface water: 10/113 Raw wastewater: 51/55
Wastewater: 148/239 oocysts River water: 38/71 oocysts
Surface water: 4/20 Wastewater and sludge: 8/36 Microscopy positive slides from river water: 10/10 Wastewater from sewage treatment plant and piggery: 6/6 River water: 11/16 C. andersoni (5), C. andersoni and C. parvum (1), W11 (1), Pig genotype II (1), C. parvum and W11 (1), C. hominis and W11 (1), C. andersoni, W11 and two new genotypes (1) Wastewater: C. hominis (78), C. parvum (16), C. meleagridis (13), C. suis (5), pig genotype II (2), mouse genotype (1), unknown (27); new genotype (1) River water: C. hominis (18), C. parvum (1), C. suis (1), pig genotype II (3), unknown (15) C. hominis (4), C. parvum (1), C. hominis and C. parvum (1) C. parvum (29), C. hominis (5), C. andersoni (1), C. muris (1), mixed (4) Surface water: C. andersoni (4), C. hominis (2), C. baileyi (1), C. meleagridis (1), C. andersoni and C. hominis (1), C. hominis and C. meleagridis (1) Wastewater: C. hominis (32), C. andersoni (13), C. parvum (8), C. muris (14), C. meleagridis (3), C. felis (3), C. canis (2), cervine genotype (9), squirrel genotype (1), W16-like (1), W19 (1), new genotype (3)
C. andersoni (2), C. baileyi (1), skunk genotype (1), C. andersoni and C. baileyi (2), C. andersoni and skunk genotype (1), all three genotypes (2) Pig genotype II (4), C. parvum + pig genotype II (1), C. parvum + cervine genotype (1)
C. parvum (12), unknown (1)
Xiao et al., 2006a
Alves et al., 2006a
Coupe et al., 2006
Hashimoto et al., 2006; Hirata and Hashimoto, 2006
Masago et al., 2006
Ryan et al., 2005a
Ruecker et al., 2005
Hanninen et al., 2005
Methods detect only C. parvum, C. hominis, C. meleagridis, and closely related parasites. CC-PCR: cell culture PCR; IMS: immunomagnetic separation. The results of the study by Nichols et al. (2006) are not included because definitive genotypes present were not determined.
SSU rRNA
IMS-nested PCR
a
COWP
IMS-PCRa
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studies have been conducted to determine the identity of Cryptosporidium oocysts in water, but results generated thus far indicate that a large proportion of Cryptosporidium oocysts in water are not from species harmful to humans (Table 5.6). Current regulatory detection methods for Cryptosporidium oocysts in water do not require species identification. Thus, the human health impact could be overestimated in risk assessment models that do not account for this. More studies with systematic sampling of different types of water are clearly needed to develop a better picture of the extent of water contamination with human-infective Cryptosporidium species in different environmental settings. Periodic determination of the species of Cryptosporidium oocysts in a watershed or source water can be helpful in developing strategies for the scientific management and protection of source water.
VII. Perspective Molecular epidemiologic studies of cryptosporidiosis are in infancy, but significant progress has been made toward better understanding of the transmission of cryptosporidiosis in humans and the public health significance of Cryptosporidium species from farm animals, companion animals, and wildlife. Cryptosporidium parvum is no longer considered a homogeneous species, and humans are known to be infected with other species in addition to C. parvum. We now have a much better appreciation of the complexity of Cryptosporidium infections in humans at species/genotype and subtype levels. We are beginning to use second-generation molecular tools to answer epidemiologic questions that are difficult to address by traditional methods, such as maintenance of immunity and cross protection, transmission dynamics in different settings, temporal and geographic distribution of Cryptosporidium genotypes and subtypes, and the role of parasite factors in the variability in transmission and clinical spectrum of cryptosporidiosis. It has become increasingly clear that the two (anthroponotic and zoonotic) cycles of cryptosporidiosis transmission previously identified (Peng et al., 1997) is an oversimplification of the complexity of cryptosporidiosis epidemiology. Increasing evidence suggests that, in certain epidemiologic settings, there can be a persistent circulation of C. parvum and other “zoonotic” species in humans without involvement of domestic animals. Thus, the contribution by humans to infections with C. parvum and other species is not known for many areas, especially areas with high C. parvum endemicity. More studies on the population genetics of C. parvum, C. hominis, and C. meleagridis, and the distribution of Cryptosporidium species and genotypes in developing countries, are required to better understand the diversity in the transmission of human cryptosporidiosis in different socioeconomic settings. The significance of the disparity in genotype distribution and segregation of certain subtypes among geographic regions is not clear. Answers to this puzzle are imperative to the understanding of the infection/contamination sources and transmission dynamics of Cryptosporidium in these areas. Likewise, the role of genotypes and subtypes in the widely reported diversity in clinical symptoms among cryptosporidiosis outbreaks requires clarification. Some biologic differences between C. hominis and C. parvum have already been identified in animal and human studies (Pereira et al., 2002; Chappell et al., 2006). Similarly, the universal distribution of certain subtypes and the identification of the involvement of particular subtypes in several major waterborne and foodborne cryptosporidiosis outbreaks indicate that certain subtypes may be more infectious and/or virulent than others. More information is needed regarding the clinical and nutritional significance of various Cryptosporidium species, genotypes, and subtypes in both children and immunocompromised patients. Cryptosporidiosis in AIDS patients causes disease ranging from brief self-limited or even asymptomatic infection to chronic diarrhea and wasting leading to death. In addition to host factors (prior exposure, status of immunosuppression, host genetic background, and antiviral treatment), parasitologic factors (species/genotypes and subtypes, virulence, infection dose, coinfection) may contribute to the variation in clinical manifestations. In industrialized countries, highly active antiretroviral therapy (HAART) has been shown to be the most effective treatment against cryptosporidiosis in AIDS patients. The use of antiretroviral drugs is increasing in developing countries. The effect of less intensive antiretroviral therapy, especially single-drug treatment, on the occurrence and pathogenesis of cryptosporidiosis in HIV+ individuals remains to be determined.
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No data are available to evaluate the role of reactivation of latent infection in cryptosporidiosis of AIDS patients, or in the person-to-person transmission of Cryptosporidium infection from mothers to infants. The relapse of cryptosporidiosis in AIDS patients who discontinued antiretroviral therapy suggests that the infection may remain in a latent stage (Maggi et al., 2000). Latent infection of C. felis can be activated by treatment of cats with prednisolone (Asahi et al., 1991), and reactivation of latent infection in pregnant animals around parturition plays an important role in the transmission of Cryptosporidium infection to newborns (Xiao et al., 1994; Ortega-Mora et al., 1999; Skerrett and Holland, 2001). Answers to these questions require collaboration among epidemiologists, clinicians, molecular biologists, and parasitologists in well-designed epidemiologic studies. In addition, comparative and histologic studies involving experimental infections of gnotobiotic pigs and human volunteers may be needed to address questions related to biologic differences among Cryptosporidium species/genotypes and subtypes. There is a clear need for more systematic investigation of outbreaks to allow for meaningful interpretation of the epidemiologic and public health significance of molecular data. Such an integrated approach will undoubtedly lead to better utilization of available molecular diagnostic tools and a better understanding of the epidemiology of cryptosporidiosis.
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IX. Appendix: Genotyping and Subtyping Cryptosporidium Oocysts in Fecal Specimens and Water Samples The following Cryptosporidium genotyping and subtyping techniques are used in the molecular epidemiology laboratory of the Division of Parasitic Diseases, Centers for Disease Control and Prevention. For fecal specimens, DNA is extracted using either the QIAamp® DNA Stool Kit (QIAGEN Inc., Valencia, CA) (Xiao et al., 2002b) or the FastDNA® SPIN Kit for Soil Samples (Q-BIOgene, Irvine, CA) (Jiang et al., 2005a). The latter requires the use of a FastPrep instrument, which may not be available in some laboratories. For environmental samples, DNA is extracted using QIAamp DNA mini kit (Qiagen) after the oocysts are isolated by immunomagnetic separation using Dynabeads® anti-Cryptosporidium kit (Dynal, Oslo, Norway) (Xiao et al., 2004c) or directly without oocyst isolation using the FastDNA® SPIN Kit for Soil Samples, with the latter technique slightly less sensitive but more economical (Jiang et al., 2005a). Cryptosporidium oocysts were first detected by nested PCR analysis of the SSU rRNA gene (Jiang et al., 2005b). Cryptosporidium species and genotypes are mostly differentiated with restriction analysis of the secondary PCR products with SspI and VspI. However, Cryptosporidium species in bovine specimens are genotyped using a combination of SspI and MboII (Feng et al., 2007), and C. muris and C. andersoni in water samples can be differentiated by the use of DdeI (Xiao et al., 2001c). Fecal specimens are generally analyzed twice with 2 µL of extracted DNA, whereas water samples are each analyzed five times because of the low number of oocysts and likely presence of mixed genotypes. With water samples, all genotyping results need confirmation by DNA sequencing (Jiang et al., 2005b). Cryptosporidium parvum and C. hominis identified are further subtyped by PCR and DNA sequencing of the GP60 gene (Alves et al., 2003b; Sulaiman et al., 2005). The two species and C. meleagridis can be further subtyped by MLST (Cama et al., 2006b; Gatei et al., 2006a, 2007). Positive (DNA of genotypes or subtypes unexpected in specimens under analysis) and negative (reagent water) are used in each PCR run.
A.
Extraction of DNA in Fecal Specimen Using the QIAamp® DNA Stool Kit
1. Store fecal specimens containing Cryptosporidium oocysts unpreserved at 4°C, or in 2.5% potassium dichromate solution at 4°C, or frozen at –20°C. 2. Transfer ~200 µL of the fecal specimen to a 2.0-mL microfuge tube. 3. Wash at least twice with distilled water by centrifugation at 1,000× g for 10 min. 4. Add 66.6 µL of 1-M KOH, and 18.6 µL of 1-M Dithiothreitol (DTT) to the pellet, and mix thoroughly by stirring with a pipette tip. 5. Incubate at 65°C for 15 min. 6. Neutralize the alkaline with 8.6 µL of 25% HCl and 160 µL of 2-M Tris-HCl (pH 8.3). Vortex the tube. 7. Add 250 µL of phenol:chloroform:isoamyl alcohol (25:24:1) solution to the tube and vortex. 8. Centrifuge at 500× g for 5 min. 9. Transfer the supernatant to a 2.0-mL microfuge tube. 10. Add 1 mL of ASL buffer from the QIAamp® DNA Stool Kit (QIAGEN Inc., Valencia, CA) to the supernatant. Incubate the mixture at 80°C for 5 min. 11. Add 1 InhibitEX tablet to each specimen and vortex immediately for 1 min or until the tablet is completely dissolved. 12. Follow the remaining procedures specified in the QIAamp® DNA Stool Kit. Elute the DNA using 200 µL of the AE Elution Buffer and store the extraction at –20°C.
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Extraction of Cryptosporidium DNA from Oocysts Isolated from Water Samples Using Immunomagnetic Separation and the QIAamp® DNA Mini Kit 1. Process 10 L of surface water or 100-L finished water samples through filtration, elution, and concentration steps, following method 1623 by the U.S. Environmental Protection Agency (EPA) (www.epa.gov/waterscience/methods/1623.pdf as of 1/1/2007). 2. Wash the packed pellet (0.5 mL of packed pellet volume or less) in 15-mL polypropylene tube twice with distilled water by centrifugation at 1,500× g for 10 min. 3. Isolate Cryptosporidium oocysts by immunomagnetic separation using the Dynabeads® antiCryptosporidium kit (Dynal A.S, Oslo, Norway) and manufacturer-suggested procedures. 4. Add 180 µL of Buffer ATL from the QIAamp DNA Mini Kit (Qiagen) to the 2.0-mL microfuge tube containing magnetic beads–Cryptosporidium oocysts (without the oocyst detachment process), and vortex for 30 s. 5. Freeze–thaw five times at –70°C (or dry ice) and 56°C. 6. Add 20 µL of proteinase K from the kit to the tube, vortex for 10 s, and incubate at 56°C overnight. 7. Add 200 µL of Buffer AL to the tube, vortex, and incubate the tube at 70°C for 10 min. 8. Follow the remaining procedures specified in the QIAamp® DNA Mini Kit. Elute the DNA using 100 µL of Buffer AE and store the extraction at –20°C.
C.
Extraction of DNA from Fecal Specimen and Water Concentrates Using the FastDNA® SPIN Soil Kit
1. Process 10 L of surface water or 100-L finished water samples through filtration, elution, and concentration steps, following method 1623 by the U.S. EPA. 2. Wash 0.5 mL of packed water concentrates or fecal suspension in 2.0-mL microfuge tubes twice with distilled water by centrifugation at 10,000× g for 3 min. 3. Aspirate the supernatant. Use the pellet immediately for DNA extraction. 4. Resuspend the washed water or fecal pellet in 978 µL of Sodium Phosphate Buffer from the FastDNA® SPIN Soil Kit (Q-BIOgene, Irvine, CA). 5. Transfer the suspension to a Lysing Matrix E Tube. 6. Add 122 µL of MT Buffer to the Lysing Matrix E Tube. 7. Secure the tube in FastPrep® Instrument (Q-BIOgene) and process for 30 s at the speed setting 5.5. 8. Centrifuge the Lysing Matrix E Tube at 10,000× g for 30 s. 9. Transfer supernatant to a clean tube. Add 250 µL of PPS reagent from the kit and mix by shaking the tube by hand 10 times. 10. Follow the remaining procedures specified in the FastDNA® SPIN Soil Kit. Elute the DNA using 100 µL of DES (DNase/pyrogen free water), and store the extraction at –20°C.
D. 1.
Genotyping Cryptosporidium by PCR-RFLP Analysis of the SSU rRNA Gene Primary PCR a. For each PCR reaction, prepare the following master mixture:
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Cryptosporidium and Cryptosporidiosis, Second Edition 10× Perkin–Elmer PCR buffer dNTP (1.25 mM) SSU-F2 primer (Table 5.1) (40 ng/µL) SSU-R2 primer (Table 5.1) (40 ng/µL) MgCl2 (25 mM) Nonacetylated bovine serum albumin (10 mg/mL) Distilled water Taq polymerase
10 µL 16 µL 2.5 µL 2.5 µL 6 µL 4 µL 56.5 µL 0.5 µL
Total
98 µL
b. Add 98 µL of the master mixture to each PCR tube. c. Add 2 µL of DNA suspension to each tube. d. Run the following PCR program: 94°C: 3 min 35 cycles of: 94ºC for 45″, 55°C for 45″, and 72°C for 1 min 72°C for 7 min 4°C soaking
2.
Secondary PCR a. For each PCR reaction, prepare the following master mixture: 10× Perkin–Elmer PCR buffer dNTP (1.25 mM) SSU-F3 primer (Table 5.1) (40 ng/µL) SSU-R4 primer (Table 5.1) (40 ng/µL) MgCl2 (25 mM) Distilled water Taq polymerase
10 µL 16 µL 5 µL 5 µL 6 µL 55.5 µL 0.5 µL
Total
98 µL
b. Add 98 µL of the master mixture to each PCR tube. c. Add 2 µL of the primary PCR reaction to each tube. d. Run the following PCR program: 94°C: 3 min 35 cycles of: 94°C for 45″, 58ºC for 45″, and 72°C for 1 min 72ºC for 7 min 4ºC soaking e. Run electrophoresis on 1.5% agarose gel with 20 µL of the PCR product, using the 100-bp ladder as the control.
3.
RFLP Analysis a. Prepare the master mixture using the following formula, which is for one restriction digestion reaction. Buffer SspI VspI DdeI (for C. muris and C. andersoni only) MboII (for bovine specimens only)
4 4 4 4
µL of New England BioLabs Buffer SspI µL of Promega Buffer D µL of New England BioLabs Buffer 3 μL of New England BioLabs Buffer 2
Water
Enzyme
22 24 24 24
4 µL 2 µL 2 µL 2 μL
µL µL µL μL
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TABLE A.1 Restriction Fragment Length Polymorphism in the SSU rRNA Gene of Common Cryptosporidium Species and Genotypes Cryptosporidium
PCR Product Size (bp)
C. hominis Rabbit genotype Monkey genotype C. parvum C. wrairi Marsupial I Opossum I Duck genotype C. bovis Deer-like genotype C. bovis in yak C. canis C. varanii Bear genotype Skunk genotype Horse genotype C. meleagridis Mouse genotype Ferret genotype C. baileyi Goose I Goose II Opossum II C. muris C. andersoni C. serpentis Cervine genotype C. suis Muskrat I Fox genotype C. felis Muskrat II C. galli Tortoise genotype Snake genotype
851 849 849 848 848 851 848 835 836 836 836 843 847 847 852 849 847 852 851 840 835 835 924 847 846 845 849 852 863 846 878 847 843 845 838
SspI Products (bp) 450, 473, 462, 450, 450, 441, 440, 432, 432, 432, 413, 417, 418, 418, 418, 450, 450, 450, 450, 573, 534, 534, 495, 449, 449, 414, 454, 454, 449, 448, 426, 417, 843 811, 804,
267, 267, 267, 267, 267, 267, 267, 267, 267, 267, 267, 267, 267, 267, 267, 267, 267, 273, 267, 267 267, 267, 429 398 397 383, 384, 378, 380, 377, 404, 375,
111, 109 109, 108, 109, 109, 107, 102, 103, 103, 103, 105, 109, 106, 110, 109, 108, 112, 111,
12, 11 11 12, 11, 34 34 34 34 34 34, 34, 34, 34, 34, 12, 11, 12, 12,
11 11
19 20 19 22 12, 11 11 11 11 11
34 34
34, 11 11, 34 21 34, 34,
14 9
14 21
34 34
VspI Products (bp) 561, 559, 559, 629, 629, 632, 629, 616, 617, 617, 617, 624, 628, 628, 460, 497, 457, 458, 458, 621, 616, 616, 668, 732, 731, 730, 461, 633, 608, 627, 659, 592, 728, 730, 619,
115, 115, 115, 115, 115, 115, 115, 115, 115, 115, 115, 115, 115, 115, 173, 133, 171, 175, 174, 115, 115, 115, 115, 115 115 115 169, 115, 115, 115, 115, 115, 115 115 115,
104, 104, 104, 104 104 104 104 104 104 104 104 104 104 104 115, 115, 115, 115, 115, 104 104 104 104,
71 71 71
104 104 104 104 104
37
115, 104 104 104, 36 104 104 104, 36
104
Note: Fragment lengths in base pairs; visible bands are shown in bold.
b. c. d. e.
Transfer 30 µL of master mixture to each tube. Add 10 µL of secondary PCR reaction to the tube and mix well. Incubate in 37°C waterbath for 2 h or overnight. Run electrophoresis on 1.2% agarose gel with the entire 40 µL of restriction digestion reaction, using the 100-bp ladder as the control. f. Identify Cryptosporidium species and genotypes based RFLP banding patterns (Table A.1).
E. 1.
Subtyping C. parvum and C. hominis by PCR-Sequencing Analysis of the GP60 Gene Primary PCR a. Preparation of master mixture. For each PCR reaction, prepare the following:
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10 µL 16 µL 5 µL 5 µL 6 µL 4 µL 51.5 µL 0.5 µL
Total
98 µL
b. Add 98 µL of the master mixture to each PCR tube. c. Add 2 µL of DNA sample to each tube. d. Run the following PCR program: 94ºC: 3 min 35 cycles of: 94°C for 45″, 50°C for 45″, and 72°C for 1 min 72ºC for 7 min 4ºC soaking
2.
Secondary PCR a. Preparation of master mixture. For each PCR reaction, prepare the following: 10× Perkin–Elmer PCR buffer dNTP (1.5 mM) GP60-F2 primer (Table A.2) (40 ng/µL) GP60-R2 primer (Table A.2) (40 ng/µL) MgCl2 (25 mM) Distilled water Taq polymerase
10 µL 16 µL 5 µL 5 µL 6 µL 54.5 µL 0.5 µL
Total
97.5 µL
b. Add 97.5 µL of the master mixture to each PCR tube. c. Add 2.5 µL of the primary PCR reaction to each tube. d. Run the following PCR program: 94ºC: 3 min 35 cycles of: 94ºC for 45″, 50ºC for 45″, and 72ºC for 1 min 72ºC for 7 min 4ºC soaking
3.
Detection of Secondary PCR Products
Run electrophoresis on 1.5% agarose gel with 15 µL of the secondary PCR product, using the 100-bp ladder as the control.
4.
DNA Sequencing a. Clean the positive secondary PCR products of the expected size using the Montage-PCR kit (Millipore, Bedford, MA). b. Store the cleaned PCR product at –20°C. c. The cleaned PCR products will be sequenced using three sequencing primers (the forward and reverse PCR primers and the intermittent primer R3 (5′-GAGATATATCTTGTTGCG-3′).
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d. For each sequencing reaction, prepare the following using components of the BigDye® Terminator V3.1 Cycle Sequencing Kit (Applied Biosystems, Foster, CA): BigDye Terminator Buffer BigDye Terminator Primer (40 ng/µL) Reagent water
4 µL 1 µL 2 µL 9.5 µL
e. Add 19.5 µL of the master mixture to a PCR tube. f. Add 0.5 µL cleaned PCR products to each tube. g. Run the following program in a PCR machine: 25 cycles of 96°C for 10 s, 50°C for 5 s, and 60°C for 4 min 1 cycle of 4°C soaking h. Clean the 20 µL of sequencing reaction products using the Centrisep Spin Column (Princeton Separations, Adelphia, NJ). i. Dry the cleaned sequencing products in a vacuum centrifuge. Store the cleaned products in a freezer. j. Add 15 µL of BigDye Formamide to dehydrated sequencing products. k. Transfer the DNA suspension to the MicroAmp Optical 96-well reaction plate (Applied Biosystems). l. Cover the plate with a 96-well Plate Septa (Applied Biosystems). m. Heat the plate in a PCR machine at 94°C for 5 min and cool the plate at –20°C for at least 3 min. n. Load the reaction product onto a 3100 AB Prism Genetic Analyzer for sequencing using proper program.
5.
Sequence Analysis a. Read out the electropherograms generated by the sequencer using the ChromasPro software (www.technelysium.com.au/ChromasPro.html) or any other software. b. Align the GP60 sequences generated with each other and reference sequences using the software ClustalX (ftp://ftp-igbmc.u-strasbg.fr/pub/ClustalX/). c. Check the sequence alignment for sequencing accuracy using the software BioEdit (www.mbio. ncsu.edu/BioEdit/bioedit.html). d. Recheck the electropherograms for any sequence uncertainty. e. Determine subtype designation based on sequence identity to reference sequences (Table A.2) and the number of trinucleotide repeats (Sulaiman et al., 2005).
F.
Multilocus Sequence Typing of C. parvum, C. hominis, and C. meleagridis
Microsatellites, minisatellites, and other polymorphic markers have been identified in the C. parvum and C. hominis genomes, and are used in multilocus sequence typing of these two species to characterize the population genetics and transmission in endemic areas (Cama et al., 2006b; Gatei et al., 2006a, 2007). Some of the primers used amplify DNA of C. meleagridis, and thus can be used effectively in subtyping this species. The sequences of these primers, the annealing temperature used, and the expected PCR product sizes are shown in Table A.2. The PCR condition used is largely the same as the GP60based subtyping shown earlier. Subtypes at each locus are determined by direct DNA sequencing of the secondary PCR products from the nested PCR, using the forward and reverse primers in the secondary PCR. Population genetic computer programs such as Arlequins (http://lgb.unige.ch/arlequin/), DnaSP (http://www.ub.es/dnasp/), and Recombination Detecting Program (RDP- http://darwin.uvigo.es /rdp/rdp.html) can be used in the analysis of multilocus sequences generated (Gatei et al., 2007).
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TABLE A.2 Primers Used in the Multilocus Sequence Typing (MLST) of C. hominis, C. parvum, and C. meleagridis Locus HSP70
GP60
CP47
CP56
Nature of Polymorphism Primer 12-bp Minisatellite
Microsatellite– TCA, TCG, TCT
Microsatellite– TAA, TGA/TAG
SNP
Sequence (5′ to 3′)
F1
ACTCTATGAAGGTATTGATT
R1 F2 R2 F1
TTAGTCGACCTCTTCAACAGTTGG CAGTTGCCATCAGTAGAG CAACAGTTGGACCATTAGATCC ATAGTCTCCGCTGTATTC
Annealing Fragment Sites Temperature (bp) 55°C 1030–1100 45°C 50°C 800–850
R1 F2 R2 F1
GGAAGGAACGATGTATCT TCCGCTGTATTCTCAGCC GCAGAGGAACCAGCATC GCTTAGATTCTGATATGGATCTAT
50°C 43°C 380–500
R1 F2 R2 F1
AGCTTACTGGTCCTGTATCAGTT ACCCCAGAAGGCGGACCAAGGTT GTATCGTGGCGTTCTGAATTATCAA CTCACAGAGTTAAAAATCACTT
55°C 50°C 660–750; not working for C. meleagridis
Mucin1
MSC6-7
MSC6-5
RPGR
63-bp Minisatellite
15-bp Minisatellite
Microsatellite– TCT/TCC
18-bp Minisatellite
TSP8 (ML2) Microsatellite– GA
R1 F2 R2 F1
GAACGCAAATATTAAGAAAAATTGAG TTGGCAATGTTGTCTTTTTCCA ATATGTAATCTGGCGCCAAAG ACTGATGTGTCAAGTGGCAATC
50°C 58°C 650–900; not working for C. meleagridis
R1 F2 R2 F1
TTACAGTTATGAGTTGCTGGT TTGATGATTCAGAATCATCTGACT GTGAGTTCTTCTTCATCTGTATAG ATTGAACAAACGCCGCAAATGTACA
R1 F2 R2 F1
CGATTATCTCAATATTGGCTGTTATTGC GCTATTTGCTATCGTCTCACATAACT CTACTGAATCTGATCTTGCATCAAGT TTGAGCCTCTTAGTATATCAAATACT
R1 F2
GAGATACCAAATGCCTTAAACCAGTATT GACTTACACCATTTCTTGGTATGCCTA 55°C
R2 F1
AACTGTCATACCACCAGTAGATGATA AAAGGTAACTCAATTGCTAAAGAT
55°C
R1 F2
TCTTCCTCTTTCTGGCTTTCAGTATT AGATCATATAGTGACACCTGATCAA
55°C
R2 F1
CCACTGAATCTTCTTTATTGTCAA TTGCAACTTTGTCAAGTA
50°C
R1 F2 R2
TATATGAGTGCACTCCAC ATTGTTTAAGTTCGGGTG AATATGGTTCCCAATGACC
55°C 55°C
545–564; not working for C. meleagridis
55°C 55°C
45°C
389–443
400–460
~340
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TABLE A.2 (CONTINUED) Primers Used in the Multilocus Sequence Typing (MLST) of C. hominis, C. parvum, and C. meleagridis Locus DZ-HRGP
ZPT
Nature of Polymorphism Primer 30-bp Minisatellite
SNP
Sequence (5′ to 3′)
F1
TGGTTGAGGTTGAAGGCCCAT
R1 F2 R2 F1
CATTTCAGCTATTTTAGCTCAACC CATTAATCTTTTAGCAAGAGTAGCTGA AATGCGTTAAGCCTTAAAGCTGG GTAAATCTTATTCAATGTACGCAA
R1
GTGATGATTTGTTTGTATTGC
F2
AGTGATTAGTAGAATCTTTGATCT
R2
ATTTGTTCAAACCACTCCATAA
Annealing Fragment Sites Temperature (bp) 55°C
590–640; not working for C. meleagridis
55°C 50°C
~ 630; not working for C. hominis and C. meleagridis
50°C
Note: The primers are based on Cama et al. (2006b) and Gatei et al. (2006a, 2007). All loci are in chromosome 6, except for HSP70, which is in chromosome 2.
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6 Diagnostics
Huw Smith
CONTENTS I. II.
Introduction .................................................................................................................................. 174 Diagnosis of Pathogenic Cryptosporidium Species..................................................................... 174 A. Cryptosporidiosis Diagnosed in Humans ....................................................................... 175 B. Cryptosporidiosis Diagnosed in Livestock and Domesticated Fowl ............................. 175 III. Oocyst Morphology...................................................................................................................... 175 A. Bright-Field, Phase Contrast (PC), and Differential Interference Contrast (DIC) Microscopy ............................................................................................ 176 B. Micrometry...................................................................................................................... 176 IV. Laboratory Diagnosis of Infection: Rationale ............................................................................. 176 V. Oocyst Concentration from Feces................................................................................................ 178 A. Biophysical Methods....................................................................................................... 178 1. Centrifugation................................................................................................... 178 2. Formol-Ether Concentration for Oocysts ........................................................ 179 Cryptosporidium Requests as Part of an Enteropathogenic 3. Parasite Screen ................................................................................................. 182 4. Oocyst Flotation............................................................................................... 182 5. Oocyst Purification........................................................................................... 183 6. Infection in the Absence of Detectable Oocysts ............................................. 183 B. Immunological Method—Immunomagnetic Separation (IMS) ..................................... 183 VI. Outbreak and Large-Scale Epidemiological Investigations......................................................... 184 A. Materials.......................................................................................................................... 184 B. Method............................................................................................................................. 184 VII. Staining Methods.......................................................................................................................... 186 A. Detection of Cryptosporidium Oocysts in Fecal Smears by Auramine Phenol ............ 186 1. Scope of Test.................................................................................................... 186 2. Preparation of Auramine Phenol (AP) ............................................................ 187 B. Detection of Cryptosporidium Oocysts Using Modified Cold Strong Ziehl–Neelsen Stain ........................................................................................................ 187 1. Scope of Test.................................................................................................... 187 VIII. Immunological Methods .............................................................................................................. 188 A. Antigen Detection Using Antibodies Labeled with Fluorescent Reporters................... 188 B. Antigen Detection Using Antibodies Labeled with Enzyme Reporters ........................ 189 1. Enzyme Immunoassays.................................................................................... 189 2. Immunochromatographic Assays..................................................................... 190 IX. Sensitivity of Detection in Feces ................................................................................................. 190 X. Antibody Detection ...................................................................................................................... 191 A. Enzyme-Linked Immunoelectrotransfer Blot (EITB) .................................................... 191 B. Enzyme-Linked Immunosorbent Assay Using Recombinant (and Other) Proteins ...... 192 XI. Biopsy........................................................................................................................................... 193
173
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XII. Molecular Diagnosis—Nucleic Acid Detection Methods ........................................................... 194 A. Extraction of Cryptosporidium DNA from Oocysts in Clinical and Environmental Samples................................................................................................... 194 1. DNA Extraction for Outbreak and Epidemiological Investigations ............... 194 2. DNA Extraction from Small Numbers of Partially Purified Oocysts............. 195 B. Primer, Gene Locus Selection, and PCR........................................................................ 197 1. 18S rRNA Gene Fragments............................................................................. 197 Cryptosporidium Oocyst Wall Protein (COWP) Gene Fragment ................... 200 2. 3. MAS-PCR ........................................................................................................ 200 4. Mixed Infections .............................................................................................. 201 5. Quantitative Real-Time PCR ........................................................................... 201 6. Typing and Subtyping for Disease and Source Tracking ............................... 202 C. Reporting Results of PCR-RFLP/Sequencing Examination .......................................... 203 XIII. Shipping of Oocysts and Oocyst DNA for Quality Assurance and Round Robin Testing ........ 203 References........................................................................................................................... 203
I.
Introduction
Although Cryptosporidium was first described in 1907, it took almost 50 years before it was recognized as a pathogen of livestock and 70 years before it was recognized as a pathogen of humans. Much of this delay was due to the lack of cost-effective methodologies to detect the parasites in clinical samples and to determine the significance of infection and disease at a population level. This chapter deals with Cryptosporidium diagnosis in clinical samples from infected human and nonhuman hosts. E.E. Tyzzer was the first person to describe Cryptosporidium in the gastric mucosa of mice (Mus musculus) in 1907. The meaning of Cryptosporidium is “hidden spore”; the word describes the transmissive stage of the parasite, namely, the oocyst. Tyzzer named the species C. muris. In 1910 he described the oocyst structure and endogenous development based on Romanowsky (Wright and Giemsa)-stained smears and histological sections of murine gastric glands. In 1912 Tyzzer reported a second species of Cryptosporidium that infected the small intestine of mice and described the morphology of the oocyst and developmental stages in histological sections and Giemsa-stained smears. This species had smaller oocysts than C. muris and was named C. parvum. There are now 16 species (Chapter 1). The size range of Cryptosporidium oocysts, which is sometimes helpful in diagnosis, extends from 3.7 × 3.0 microns (C. scophthalmi) to 8.0 × 6.4 microns (C. galli) (Chapter 1). Based on histological and microscopic findings, Cryptosporidium was recognized as a cause of morbidity and mortality in young turkeys in 1955 and as a cause of scours (diarrhea) in calves in 1971, but it was not until 1976 that the first two human cases of cryptosporidiosis were described histologically (Nime, et al., 1976; Meisel et al., 1976; Chapters 1 and 8). Between 1978 and 1980, the Giemsa stain was used to detect oocysts in smears of cattle and human feces (Pohlenz et al., 1978; Tzipori et al., 1980). In 1981 a modified Ziehl–Neelsen technique was used to stain oocysts (Henricksen and Pohlenz, 1981). In 1982 cryptosporidiosis became associated with significant mortality in AIDS patients (Pitlik et al., 1983), and in the same year the first cryptosporidiosis outbreak was recorded in immunologically healthy persons. The development of noninvasive laboratory-based methods for staining oocysts in feces led to widespread investigations of occurrence and to the awareness that Cryptosporidium is a significant cause of diarrhea in immunocompetent and immunocompromised hosts, worldwide.
II.
Diagnosis of Pathogenic Cryptosporidium Species
Species definition and identification in the genus Cryptosporidium is constantly changing, with the addition of new species based on molecular criteria and supported by biological and morphometric data. In human and nonhuman hosts, molecular methods such as the polymerase chain reaction (PCR), restriction fragment length polymorphism (RFLP), and DNA sequencing have demonstrated a greater number of Cryptosporidium species than previously thought (Chapter 5). Currently, there are 16 valid
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species and over 40 genotypes, some of which eventually might represent different species (Xiao et al., 2004; Chapter 1).
A.
Cryptosporidiosis Diagnosed in Humans
Cryptosporidium parvum, C. hominis, C. meleagridis, C. felis, C. canis, C. suis, C. muris, and C. andersoni can infect humans, but C. parvum and C. hominis are the most frequently identified species (Xiao et al., 2004; Cacciò et al., 2005; Palmer et al., 2003; Morse et al., 2007; Chapters 1, 4, and 5). Cryptosporidium parvum is the species normally, but not exclusively, associated with zoonotic transmission. There are a few reports of C. hominis infections in cattle and sheep. Cryptosporidium hominis monkey genotype and Cryptosporidium cervine genotype have been described in humans also. In humans, the incubation period (the time from the ingestion of oocysts to onset of clinical disease) can range from 2 to 14 days, although 3 to 8 days is usual. Gastrointestinal symptoms usually last from 3 to 21 days, possibly up to 6 weeks, and persistent weakness, lethargy, mild abdominal pain and bowel looseness might persist for a month (Casemore, 1987; Chapter 8). Oocyst shedding can be intermittent and can continue for up to 50 days after the cessation of symptoms (mean of 7 days).
B.
Cryptosporidiosis Diagnosed in Livestock and Domesticated Fowl
Cryptosporidium parvum and C. andersoni have been associated with morbidity in livestock, whereas C. baileyi, C. galli, and C. meleagridis have been associated with morbidity in domesticated fowl (see Chapters 15 and 18). The diagnostic difficulty arises when microscopy is employed and oocysts cannot be distinguished morphologically at the species level. Cryptosporidium parvum causes diarrhea primarily in young, preweaned ruminants; postweaned and adult animals occasionally become infected and oocysts can be detected in apparently healthy as well as sick animals (Chapters 1 and 18). Diarrheic calves can excrete between 5 × 105 to 2 × 106 oocysts per gram of feces. Cryptosporidium andersoni infects the abomasum (stomach) of older calves and mature cattle. It does not cause diarrhea but is thought to reduce weight gain and milk yield. Infected animals can excrete oocysts for months but tend to excrete fewer oocysts (300 to 1500 oocysts per gram of feces) than those infected with C. parvum (Enemark et al., 2002). Cryptosporidium baileyi, C. meleagridis, and C. galli infect domesticated fowl. Other unnamed Cryptosporidium spp. infect ostriches, quail, and other birds (Chapter 15). Cryptosporidium baileyi infects the epithelium of the bursa of Fabricius and cloaca of chickens; the trachea and the conjunctiva are less frequent sites, although respiratory cryptosporidiosis can cause severe morbidity in chickens and turkeys. Cryptosporidium meleagridis, characterized histologically by villous atrophy, crypt hyperplasia, and shortened microvilli in the ileum of turkeys, causes severe diarrhea in poults (Current, 1997). Cryptosporidium galli develops in the epithelium of the proventriculus, causing disease in chickens and finches. An inadequately described isolate of Cryptosporidium with oocysts smaller than those of C. baileyi was associated with respiratory and intestinal infections in commercial quail but was not infectious for chicken or turkeys (Current, 1997).
III. Oocyst Morphology Sporulated oocysts are smooth, thick-walled, colorless, spherical, or slightly ovoid bodies, containing four elongated sporozoites and a residual body. Contents are difficult to observe by light microscopy. Most oocysts are similar in shape and overlap in size (Chapter 1). Their morphometry can be helpful in distinguishing oocysts from other microscopic objects but generally is not useful for determining species. Because oocysts of C. muris, C. andersoni, and C. galli are larger than oocysts of species that infect the lower gastrointestinal tract, their size is helpful in differentiating gastric from intestinal Cryptosporidium spp.
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Oocysts can be detected by bright-field, phase-contrast, or differential interference contrast microscopy. To obtain accurate measurements, oocysts must be suspended in a medium, such as water, that does not cause their occlusion, expansion, contraction, or distortion. Because they are small, higher-magnification objectives must be used to confirm their presence. Measurements should be taken with the 100× objective. These methods generally are not advocated for detecting oocysts in feces that have not been subjected to concentration, but can be used for detecting, enumerating, and measuring purified or partially purified oocysts.
A.
Bright-Field, Phase Contrast (PC), and Differential Interference Contrast (DIC) Microscopy
Even at magnifications of 400 to 1000×, many structural details in the colorless oocysts are undetectable by light microscopy. PC and DIC optics can enhance the contrast through a phase shift of light, reducing brightness based on refractive indices. DIC microscopy provides a sharp image of the oocyst perimeter for measurement and a clearer image of sporozoites and the residual body (Figure 6.1). Detection can be enhanced by stains, but because many staining methods involve air drying and fixation of wet smears, oocysts become distorted and consequently do not provide accurate morphometric data. Oocysts can be stained in suspension using immunofluorescent methods and measured with reasonable accuracy.
B.
Micrometry
Measurement of the size and shape of Cryptosporidium oocysts ensures that their morphometrics fall within the accepted range of standard parameters for the species. The standard unit of measurement is the micron (µ = 0.001 mm). Measurements are made using a stage micrometer in conjunction with an eyepiece micrometer. The stage micrometer is a glass slide containing a millimeter scale graduated in microns. The eyepiece micrometer is a disk of glass or plastic bearing a graduated scale subdivided into millimeters. For each objective lens determine the number of divisions in the eyepiece micrometer that correspond to a definite number of divisions on the stage micrometer. Divide the latter by the former to calculate the length in microns of each division in the eyepiece micrometer. For measuring, superimpose the eyepiece micrometer over the oocyst, and count the number of divisions for length and width. Translate the number of divisions into microns.
IV.
Laboratory Diagnosis of Infection: Rationale
Numerous techniques, including histology and ultrastructural examination of biopsy material for lifecycle stages (Chapter 1), examination of feces for the presence of oocysts, and detection of Cryptosporidium antigens and DNA, have been used to diagnose infection in humans and animals. Molecularbased techniques are required for species identification (Chapters 1 and 5). Many fluids and tissues can be submitted for analysis, including stools, sputum, bile, mucoid secretions, and tissue biopsies, but stools are the primary specimens examined for enteropathogenic species and genotypes. All specimens are amenable to staining, antigen detection/localization, and molecular-based methods. Cryptosporidium is a Biosafety Level II organism, and laboratory procedures must be conducted in a microbiological safety cabinet. Clinical specimens might contain other pathogens and should be processed in accordance with local safety codes. Diagnosis is based primarily on demonstration of oocysts in unpreserved or preserved stools (Table 6.1). Fecal specimens can be preserved in 10% formalin, sodium acetate-acetic acid-formalin (SAF), and polyvinyl alcohol (PVA) fixatives. Many oocyst-staining methods cannot be performed on stools preserved in PVA. Oocyst viability is retained following storage in 2.5% potassium dichromate or at 4°C, but storage in formalin for extended periods should be avoided if molecular analyses are to be performed.
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FIGURE 6.1 (A color version of this figure follows page 242.) Panel of oocyst images. (A) Two C. parvum oocysts following cesium chloride purification and photographed in suspension using Nomarski differential interference contrast microscopy. Note the presence of three of the four sporozoites in the upper oocyst and the typical padlock arrangement of sporozoites in the lower oocyst. Magnification 1250×. (B) A group of C. parvum oocysts prepared from an air-dried formol ether concentrate of stool and stained with the modified Ziehl–Neelsen acid-fast stain. Cryptosporidium oocysts stain red and appear as spherules. Note the variation in staining intensity of individual oocysts in this image. The contents of many oocysts will appear amorphous, but some contain the characteristic crescentic forms of sporozoites. Magnification 1250 ×. (C) A group of C. parvum oocysts prepared from an air-dried formol ether concentrate of stool and stained with the auramine phenol stain. Cryptosporidium oocysts appear ring shaped and exhibit a characteristically bright green fluorescence against a dark background. Magnification 1250×. (D) A group of C. parvum oocysts prepared from an air-dried formol ether concentrate of stool and stained with a commercially available fluorescein isothiocyanate-labeled monoclonal antibody reactive with exposed epitopes on Cryptosporidium spp. oocysts. Note the following three characteristics of Cryptosporidium spp. oocysts using this stain: (a) characteristic apple-green fluorescence delineating the oocyst wall, under the FITC filter set. Often the fluorescence has an increased intensity around the entire circumference of the oocyst, with no visible breaks in oocyst wall staining; (b) round or slightly ovoid objects; (c) a size of 4 to 6 µm in diameter for most human pathogens. Magnification 1250×.
Stool samples from most symptomatic cases contain large numbers of oocysts, and if the specific request is for Cryptosporidium detection, most laboratories will process the specimen as a direct smear. Standard staining and immunological techniques should result in a positive diagnosis. Soft, unformed, and diarrheic stools should be mixed thoroughly, a sample removed with a pipette or a swab, and thinly smeared onto a microscope slide. Formed stools should be comminuted into a slurry in 0.9% (150-mM)
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TABLE 6.1 Some Differences Among Species Infecting Humans, Livestock, and Domesticated Fowl Within the Genus Cryptosporidium Oocyst Dimensions (µm)
Species
Site of Infection
C. hominis C. parvum
4.5 × 5.5 4.5 × 5.5
Small intestine Small intestine
C. C. C. C. C. C. C. C. C.
4.41 × 5.05 4.5 × 5.0 3.68–5.88 × 3.68–5.88 4.5–4.0 × 4.6–5.2 5.5 × 7.4 5.6 × 7.4 (5.0–6.5 × 8.1–6.0) 4.0–5.0 × 4.8–5.6 4.17–4.76 × 4.76–5.35 4.6 × 6.2
Small intestine Small intestine Small intestine Intestine Stomach Stomach Small intestine Small intestine Trachea, bursa of Fabricius, cloaca Proventriculus
suis felis canis meleagridis muris andersoni wrairi bovis baileyi
C. galli
6.2–6.4 × 8.0–8.5
Major Host
Infectious to Humans
Humans Neonatal mammalian livestock, humans Pigs Cats Dogs Turkeys Rodents Cattle Guinea pigs Cattle Poultry
X X X X
Finches, chicken
X
saline to produce a more liquid emulsion before the smear is made. The best smear thickness is achieved when either the hands of a watch or the print on a page can just be seen through the preparation. Smears should be air-dried, avoiding excessive heat that can distort oocysts, and then stained. Oocysts are more readily detected in concentrates from watery specimens than from formed stools. In specimens containing small numbers of oocysts, increased sensitivity can be achieved by employing a concentration method followed by staining.
V. A.
Oocyst Concentration from Feces Biophysical Methods
Methods to concentrate oocysts from feces while reducing debris increase the sensitivity of diagnostic methods (Garcia et al., 1983). The most common concentration methods include Sheather’s (1923) sucrose flotation method and formol-ether (formol-ethyl acetate) concentration. Although some investigators find Sheather’s to be the superior method (Ma and Soave, 1983), it is rarely used in analysis of human stools because of their fat content. If oocysts are not thoroughly washed, sucrose can also interfere with staining and adherence to microscope slides (Casemore, 1985). The modified formol-ether method, reported to be more sensitive than the method of Allen and Ridley (1970), is recommended for stools containing few oocysts (e.g., follow-up specimens from individuals who have recovered; Casemore et al., 1985) but other conventional fecal flotation methods can be used for animal feces. Low oocyst excretors (e.g., asymptomatic excretors) probably will not be diagnosed, because oocyst numbers will be below the limit of detection of these conventional methods.
1.
Centrifugation
Oocysts settle more rapidly if the stool suspension is subjected to centrifugation. However, partly digested food particles will also sediment and can mask the presence of oocysts in microscopic slide preparations. To overcome this potential problem, remove larger particles before centrifugation by filtering emulsified stool through a sieve. Several sieving options are available such as folded medical gauze, which is then disposed of, kitchen-type mesh strainers, and metal or plastic screened sieves of multiple pore sizes. The quantity of feces examined can influence the findings. For example, when 25 replicates of bovine feces were each spiked with 10, 50, and 100 oocysts of C. parvum per gram of feces and subjected to sieving followed by cesium chloride density gradient centrifugation, more positive detections at each
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spiking level were obtained from 15-g samples than from 10- or 5-g samples (Fayer, personal communication). For human stools, after adding formalin for fixation and preservation, the level of detection can be increased by adding ether or ethyl acetate to remove fats and oils. After centrifugation, a fatty plug can be seen at the interface of the two liquids. The ether layer, fatty plug, and formalin layer are discarded and the pellet retained for examination. Many modifications to this procedure have been advocated. The following protocol modified from Allen and Ridley (1970) achieves a concentration of 15- to 50-fold.
2. a.
Formol-Ether Concentration for Oocysts Materials
Disposable gloves, 15-mL conical glass centrifuge tubes, disposable wooden applicator sticks, sieve (425-µm aperture size, 38-mm diameter*), 50-mL Pyrex beaker**, centrifuge with 15-mL swing-out buckets, glass microscope slides, coverslips, diamond marker, bright-field microscope with 10× and 40× objective lenses, 10% formalin, and diethyl ether (or ethyl acetate).
b.
Method 1. Wear gloves. Sample approximately 0.5- to 1-g feces with an applicator stick*** and place in a clean centrifuge tube containing 7 mL of 10% formalin. If the stool is liquid, dispense about 750 µL into the centrifuge tube. 2. Break up the sample thoroughly and comminute with an applicator stick. 3. Filter the resulting suspension through a sieve or gauze into a beaker, and pour the filtrate back into the same tube.*,**** 4. Add 3 mL of diethyl ether (or ethyl acetate*****) to the formalinized solution, seal the neck of the tube with a rubber bung (or a gloved thumb over the top of the tube), and shake the solution vigorously for 30 s. Invert the tube a few times during this procedure, and release the pressure developed gently by removing the rubber bung (or your thumb) slowly. 5. Centrifuge the tube at 1200****** × g for 2 min. 6. Loosen the fatty plug with a wooden stick by passing the stick between the inner walls of the tube and the plug. Discard the plug and the fluid both above and below it by inverting the tube, allowing only the last one or two drops to fall back into the tube. Discard this fluid containing diethyl ether and formalin into the marked resealable liquid waste container kept in the fume cupboard. Resuspend the pellet by agitation. 7. Pour the whole, or the majority of the resuspended pellet onto a microscope slide, or transfer the resuspended contents onto a microscope slide with a Pasteur pipette. Allow to air-dry. 8. Stain for Cryptosporidium sp. oocysts using the protocols identified in Sections VIIA and B or a commercially available kit for the detection of oocysts on microscope slides by immunofluorescence (see Table 6.2 and Section VIIIA).
* 425-µm aperture size, 38-mm diameter is equivalent to 36 mesh British Standard (BS 410-86) or 40 mesh American Standard (ASTM E11-81). ** The skirt of the sieve should fit neatly into the rim of the beaker. *** The sample should include portions from the surface and from within a formed stool. **** Debris trapped on the sieve is discarded. Both the sieve and the beaker should be washed thoroughly in running tap water between each sample. ***** Ethyl acetate and diethyl ether are flammable and should be used in well ventilated areas, away from flames. There is no significant difference in the number of recovered oocysts or their viability when using either formalin ether or formalin ethyl acetate. Diethyl ether is more effective at extracting fats from stool samples than ethyl acetate, and yields cleaner pellets. ****** If the Cryptosporidium request is part of an enteropathogenic parasite screen (Section VA3), centrifugation at
speeds higher than 750 × g for > 2 min is not advised because some helminth ova rupture and collapse at high centrifugal forces.
DFAT
Strategic Diagnostics Inc.
Cryptosporidium Antigen Detection Microwell ELISA RIDASCREEN® Cryptosporidium Color Vue Cryptosporidium CRYPTOSPORIDIUM II SafePath® Cryptosporidium Immunoassay Kit 96-well EIA Cryptosporidium Elisa ColorPac™ Giardia/ Cryptosporidium Rapid Assay Test Kit Crypto-Strip Quick Test (Dispstick) Cryptosporidium ImmunoCard STAT!® Crypto/Giardia
http://www.meridianbioscience.com Feces IC
Meridian Diagnostics Inc.
http://www.corisbio.com/ http://www.diagnostics.be/ Feces Feces IC IC
Coris BioConcept Dispstick Cypress Diagnostics
http://www.pointe.com.pl/produkty_testy_weterynatyjne.html http://www.bd.com/ds/productCenter/DT-ColorPacGC.asp
EIA-plate IC
Pointe Scientific (Poland) Becton Dickenson
www.r-biopharm.com/clinical/products_cryptosporidium http://www.alexon-trend.com/ http://www.techlab.com/ http://www.safepath.com/crypto.html
http://www.virology.net/garryfavwebvirlabs.html
http://www.alexon-trend.com/ http://www.oxoid.com http://www.cellabs.com.au/ www.tcsbiosciences.co.uk http://www.dako.com/ http://www.rapidtest.com http://www.jnii-usa-bharat.com/
http://www.sdix.com
http://www.waterborneinc.com/
http://www.btfbio.com/ www.tcsbiosciences.co.uk / www.tcswatersciences.co.uk http://www.cellabs.com.au/ www.tcsbiosciences.co.uk / www.tcswatersciences.co.uk http://www.meridianbioscience.com
Uniform Resource Locator (URL)
Feces Feces
Feces Feces Feces
EIA-plate EIA-plate EIA-plate
r-biopharm Seradyn Inc. TechLab SafePath
Feces Feces Feces Feces
EIA-plate EIA-plate EIA-plate
Feces
EIA-plate
Dako Corp. Diagnostic Automation Inc. JN-International Medical Corp. LMD Laboratories
EIA-plate
EIA-plate
Feces, foods, environmental Feces, foods, environmental Feces, foods, environmental Feces, foods, environmental Feces, foods, environmental Feces
Sample Type
180
Crytpo CELISA, Crypto/Giardia CELISA. IDEIA Cryptosporidium Cryptosporidium ELISA Cryptosporidium EIA
Alexon Incorporated Oxoid Ltd. (ThermoFisher) Cellabs Pty, Ltd.
DFAT
Waterborne Inc.
ProSpecT/Cryptosporidium
DFAT
Meridian Diagnostics Inc.
MeriFluor™ Cryptosporidium/Giardia Crypt-a-Glo AquaGlo G/C HYDROFLUOR™ Combo-II
DFAT
DFAT
BTF Precise Microbiology Pty, Ltd. Cellabs Pty, Ltd.
EasyStain Crypto, EasyStain Crypto/Giardia. Crypto-Cel and Crypto/Giardia-Cel.
Type of Test
Manufacturer/Distributor
Kit Name
Some Examples of Commercially Available Diagnostic Kits for the Detection of Cryptosporidium
TABLE 6.2
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IMS IMS
IMS
Waterborne Inc.
Aureon Biosystems GmbH
IC IC IC IMS
Promar Labs. r-biopharm Biosite Inc. TCS Water Sciences Invitrogen
IC
Oxoid
Feces, foods, environmental
Feces Feces Feces Feces, foods, environmental Feces, foods, environmental Feces, foods, environmental
Feces
http://www.aureonbio.com/
http://www.waterborneinc.com/
http://www.invitrogen.com/
http://www.promarlabs.com/Test.cfm http://www.r-biopharm.com/general/main.php http://www.biosite.com/products/parasite.aspx http://www.tcswatersciences.co.uk/
http://www.oxoid.com/
Note: DFAT = direct immunofluorescence antibody test; EIA-plate = enzyme-linked immunosorbent assay (ELISA); IC = immunochromatography (lateral flow); IMS = immunomagnetic separation.
Dynabeads® anti-Cryptosporidium Kit IMS–Grab™ Beads Cryptosporidium oocysts, Giardia cyst Crypto Kit, Giardia Kit, and Crypto/Giardia Kit
Xpect* Giardia/Cryptosporidium Test Kit Dipstick Dipstick Test RIDA® Quick Cryptosporidium Triage® Parasite Panel Isolate® Cryptosporidium IMS
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A commercial device is available for concentrating helminth ova, larvae, and protozoan cysts and oocysts according to a modification of the formalin-ether method of Ritchie. Sold as the Fecal Parasite Concentrator (FPC, Evergreen Scientific, Los Angeles, California) it is an enclosed system, and consists of two polypropylene tubes, a flat-bottomed tube used for emulsifying the stool, and a conical tube used for centrifugation, with an interconnecting sieve. The method states that both fresh and preserved (10% formalin, merthiolate-iodine-formalin, polyvinyl alcohol, and sodium acetate-formalin) stool specimens can be used. Water ether concentration (where grade 1 water [deionized water that has undergone reverse osmosis; conductivity 0.0 to 0.09 µS/cm; British standard No. 3978] or similar is substituted for formalin) can be used when further molecular analyses are required. A greater volume (up to 1 mL) of watery feces than formed (up to 1 g) can be purified using water ether concentration. Water ether concentration recovers higher percentages of C. parvum oocysts than either sucrose or zinc sulfate flotation. Water ether concentration recovers 46.4 to 74.9%, sucrose flotation recovers 24 to 64.8%, and zinc sulfate flotation recovers 21.6 to 40.9% (Bukhari and Smith, 1995).
3.
Cryptosporidium Requests as Part of an Enteropathogenic Parasite Screen
When stool samples are to be processed as direct smears, specialized parasitology diagnostic and reference laboratories might be requested to investigate a sample for the presence of a variety of parasitic enteropathogens, including Cryptosporidium. In the absence of fresh or preserved, recently voided stool samples, in which the vegetative and transmissive stages of enteropathogenic protozoa can be sought, most specialized laboratories will use the formol ether (ethyl acetate) method for enteropathogenic parasite screens. Cryptosporidium can either be detected separately from other parasitic enteropathogens by staining direct smears, or as a component part of the enteropathogenic parasites screen. In the author’s laboratory, the transmissive stages of parasitic enteropathogens are sought in wet films following formol ether concentration, then the coverslip is removed, and the wet film is air-dried and subjected to auramine phenol staining (Section VIIA). Modified Ziehl Neelsen or a commercially available kit for the detection of oocysts on microscope slides by immunofluorescence (Table 6.2 and Section VIIIA) can also be used. Before removing the coverslip for Cryptosporidium staining, the wet film can be analyzed by fluorescence microscopy for Cyclospora and Isospora oocysts that autofluoresce a sky blue color under UV light.
4.
Oocyst Flotation
The density of intact, viable C. parvum oocysts is approximately 1.05 g/m3 (settling velocity = 0.0018 m/h) (Anon., 1990), and the flotation principle utilizes a liquid suspending medium that is denser than the oocysts to be concentrated. When feces are mixed with the flotation solution, oocysts rise to the surface and can be skimmed or aspirated from the surface. For a flotation fluid to be useful in diagnostics, it must not only have a greater specific gravity than the oocysts, but must not damage or render them unrecognizable. Brine, a concentrated aqueous NaCl solution, has a specific gravity of 1.12 to 1.20, and specimens should be examined within 5 to 20 min after flotation. Sucrose solutions induce less oocyst distortion, although prolonged storage in sucrose can result in morphological changes (Robertson et al., 1993b). The oocyst flotation techniques described later selectively concentrate viable oocysts (Bukhari and Smith, 1995). Oocysts in liquid stools concentrate better than those in semiformed and formed stools because fewer are attached to large fecal particulates. Semiformed and formed stools should be thoroughly mixed in grade 1 water or brine to ensure maximal separation of oocysts from fecal debris. Oocysts can be partially purified and concentrated by the formol ether or water ether concentration methods before being concentrated by flotation methods.
a.
Sucrose Flotation Protocol
Sucrose solution (specific gravity 1.18) is prepared in a glass beaker by adding 256 g of sucrose to 300 mL of grade 1 water. The solution is gently heated (< 60°C) and continuously stirred with the aid of a magnetic stirrer on a hot plate stirrer, until the sucrose has dissolved completely. The sucrose solution can be stored at 4°C until used.
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Fecal samples are comminuted in 10 mL of water, then underlayed with an equal volume of cold sucrose solution and centrifuged (1050 × g, 15 min). Following centrifugation, 10 mL of fluid from the sucrose–water interface is withdrawn using a 10-mL syringe and a hypodermic needle and transferred in to a clean 50-mL centrifuge tube and washed thrice in grade 1 water.
b.
Saturated Salt Flotation Protocol
Saturated salt solution (specific gravity 1.2) is prepared by adding approximately 200 g of sodium chloride to 200 mL of grade 1 water. The solution is gently heated (< 60°C) while being continuously stirred and while small amounts of sodium chloride (approximately 10 g) are added at 10-min intervals until the solution becomes saturated. The saturated salt solution is then held at 4°C. The cold saturated salt solution is then transferred into a 500-mL measuring cylinder and, where necessary, its specific gravity is adjusted to 1.2 by adding cold grade 1 water (4°C). The saturated salt solution is transferred in to a screw cap glass bottle and stored at 4°C until used. Fecal samples are thoroughly mixed in 10-mL brine (specific gravity1.2) and centrifuged (1050 × g, 2 min). Following centrifugation, 2 mL of fluid from the meniscus (containing the oocysts) is removed, washed thrice in grade 1 water and resuspended to the required volume.
5.
Oocyst Purification
Purified oocysts are required for basic and applied research, animal and human infectivity studies, disinfection and water treatment studies, and other purposes. Commercial supplies of purified C. parvum oocysts are available in North America, the United Kingdom, and Australia and can be shipped under International Air Transport Association (IATA) regulations. Cryptosporidium muris and C. baileyi oocysts can also be supplied commercially, but demand is far lower than for C. parvum. Most purification procedures involve removal of fecal debris by sieving through a series of graded sieves, the removal of fecal fats and lipids, and oocyst concentration. Oocysts can be purified in the laboratory using methods based on sucrose and salt flotation (described earlier), discontinuous sucrose and isopycnic Percoll gradients (Arrowood and Sterling, 1987), cesium chloride and Percoll (Kilani and Sekla, 1987), discontinuous sucrose and cesium chloride (Arrowood and Donaldson, 1996), acid flocculation and sodium dodecyl sulfate suspension (Hill et al., 1990), water or formol ether (Bukhari and Smith, 1995; Nichols et al., 2006a) and immunomagnetic separation (described later).
6.
Infection in the Absence of Detectable Oocysts
Oocysts might not be detectable in clinical samples from all cryptosporidiosis cases, and the absence of oocysts in repeated submissions of samples from symptomatic individuals does not necessarily indicate the absence of infection. Oocyst abundance can be low in recuperating immunocompetent cases, and in immunosuppressed individuals who have sufficient immunity to downregulate oocyst production, but insufficient immunity to down regulate asexual reproduction in enterocytes (e.g., transplant patients, patients with deficient cell-mediated immunity, primary immunodeficiency diseases, primarily singlegene disorders of the immune system; McLauchlin et al., 2003). In these instances, and particularly when clinical suspicion is high, oocyst-negative stool samples should be subjected to antigen and PCR-based detection, because sufficient Cryptosporidium antigen or DNA from asexual life-cycle forms can be present in feces. For PCR-based methods, nested PCR methods, being more sensitive than direct PCR methods, are likely to have a higher diagnostic index.
B.
Immunological Method—Immunomagnetic Separation (IMS)
IMS is not normally used for routine clinical or veterinary diagnosis, because sufficient oocysts are present in either unconcentrated stools or stools concentrated using biophysical methods (Section Va). IMS is limited by time and cost constraints. It is an option when investigating specific cases of infection or disease, particularly when the index of clinical or epidemiological suspicion is high.
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The principle underpinning IMS is that surface-exposed oocyst epitopes are bound to magnetizable beads coated with monoclonal antibodies (mAbs) recognizing oocyst wall protein. The bead–oocyst complex is concentrated in suspension by applying magnetic force from a permanent magnet that attracts the bead–oocyst complex to the side of a test tube. Once attracted to the magnet, oocysts are held in place, and the suspending fluid is removed by aspiration. Bead–oocyst complexes can be dissociated in an acidic solution, liberating concentrated and purified oocysts (Campbell and Smith, 1997). Both paramagnetic colloidal magnetite particles (40 nm) and iron-cored latex beads (e.g., Dynal® beads) have been used to concentrate oocysts. Antibody-coated paramagnetic colloidal magnetite particles have the advantages that (1) their small size is beyond the resolving power of the light microscope lens and does not interfere with oocyst and cyst identification by phase contrast microscopy, and (2) the surface-area-to-volume ratio of these particles, being larger than that for larger particles, can allow more antibody-binding sites per particle, theoretically producing a more reactive particle. Cryptosporidium species, genotype and isolate, antibody affinity and isotype, oocyst epitope expression density, and the physicochemical environment can affect epitope–paratope interaction and, thus, the IMS performance. Epitope–paratope interaction is most stable with high-affinity antibodies. Many commercially available Cryptosporidium mAbs used in IMS kits are IgM isotype (Table 6.3), but some use an isotype-switched, affinity-matured IgG (e.g., IgG3). Affinity-matured paratopes normally bind epitopes tighter than nonisotype-switched paratopes. In IMS procedures, antibody capture and release are tradeoffs, in that paratopes that bind strongly to their epitopes are less easy to dissociate. Lower-affinity paratopes can bind oocyst epitopes effectively in favorable conditions, but less effectively in adverse conditions. Higher-affinity mAbs bind oocyst epitopes in more adverse conditions. Therefore, the local environment can have a major influence on IMS performance with factors such as pH, turbidity, and divalent cations affecting antibody binding. Commercial IMS kits, primarily based on iron-cored beads, are available (Table 6.2). Using an IMS procedure, a mean of 63.0 ± 8.7 (n = 19) oocysts were recovered from oocyst-negative stool samples seeded with 100 ± 2 flow cytometry sorted C. parvum oocysts (Nichols et al., 2006a). Recovery efficiencies vary according to fecal consistency: liquid (49.8 ± 8%; n = 5), semisolid (43.2 ± 6.4%; n = 5), and solid feces (29.5 ± 4.5%; n = 5). IMS can also selectively concentrate oocysts from samples with a low abundance of oocysts for PCRbased analyses.
VI. Outbreak and Large-Scale Epidemiological Investigations Often, rapid methods are required for outbreak investigations because of the large number of samples and the requirement for timely confirmation of infection and subsequent typing and subtyping investigations. For analysis in large-scale epidemiological investigations, a small-scale, rapid formol etherbased method was developed (Nichols et al., 2006b).
A.
Materials
Disposable gloves, 1.5-mL plastic centrifuge tubes, disposable wooden applicator sticks, microcentrifuge with 1.5-mL buckets, glass microscope slides, coverslips, diamond marker, bright-field and/or fluorescence microscope with 10× and 40× objective lenses, 10% formalin, and diethyl ether (or ethyl acetate).
B.
Method 1. Wear gloves. Add 500-mg formed stool feces with an applicator stick or 200-μL liquid stool into each 1.5-mL microcentrifuge tube and comminute with 700 μL of grade 1 water. 2. Add approximately 300 μL of ether (or ethyl acetate) to each suspension, cap each tube, and vortex the capped tubes for 30 s. 3. Centrifuge the capped microcentrifuge tubes at 14,000 × g for 1 min.
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TABLE 6.3 Range of Commercially Available Antibodies to Cryptosporidium Manufacturer/Distributor: AbD Serotec (http://www.serotec.com/asp/index.html) Monoclonal Antibody Polyclonal Specificity (isotype/subtype) Antibody Unconjugated Mouse Mouse Mouse Mouse Mouse Mouse
anti-Cryptosporidium antihuman Cryptosporidium antibovine Cryptosporidium anti-Cryptosporidium anti-C. parvum (oocysts) anti-C. parvum (oocysts)
Clone BGN/4H5 (IgA) Clone BGN/3E8 (IgM) Clone BEL 0126 (IgG3) Clone BEL 0126 (IgG3) (IgG3) Clone 18.280 (IgM)
Conjugated
FITC
Manufacturer/Distributor: Biodesign International (http://www.biodesign.com/) Goat anti-Cryptosporidium Mouse anti- C. parvum Mouse anti-C. parvum (fecal oocysts) Mouse anti-C. parvum (intact oocysts) Mouse anti-C. parvum (50-kDa inner oocyst wall antigen) Mouse anti-C. parvum (15-kDa sporozoite surface glycoprotein)
Clone Clone Clone Clone
BEL0126 (IgG3) 107(IgG3) BDI370 (IgG) 2D7 (IgM)
v
Clone 2B3 (IgG2b)
Manufacturer/Distributor: Chemicon (http://www.chemicon.com/) Anti-Cryptosporidium Anti-C. parvum
Clone 18.280 (IgM) Clone MAB-C1 (IgM) Manufacturer/Distributor: GeneTex (http://www.genetex.com/)
Goat anti-Cryptosporidium Mouse anti-C. parvum
Clone 107 (IgG3)
Manufacturer/Distributor: Novus Biologicals (http://www.novusbio.com/home/index) Goat anti-Cryptosporidium Mouse anti-C. parvum
Clone 107 (IgG3)
Manufacturer/Distributor: ViroStat Inc. (http://www.virostat-inc.com/) Anti-C. parvum (oocysts) Cat. No. 7601 Anti-C. parvum (oocysts) Cat. No. 7631
(IgG3) (IgG1)
Manufacturer/Distributor: Vision BioSystems (http://www.vision-bio.com/) Anti-Cryptosporidium
4. Retain the sediment, but aspirate the fatty plug and the fluid both above and below the plug to waste. 5. Wash the sediment three times with 1 mL of grade 1 water each time, cap the tubes then vortex (30 s). Centrifuge (14,000 × g for 1 min), then aspirate the supernatant to waste. The partially purified sample can then be subjected to staining, antigen detection, and/or PCR-based methods. The sample is ready for PCR applications once the sediment is washed in lysis buffer (50-mM Tris-HCl pH 8.5, 1-mM EDTA, 0.5% SDS), and the DNA is extracted by freeze thawing (Section XIIA2; Nichols et al., 2006b).
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VII. Staining Methods Tyzzer (1910, 1912) used Giemsa to stain Cryptosporidium life-cycle stages in murine gut mucosal smears. Nearly 70 years later, Pohlenz et al. (1978) used Giemsa stain to detect oocysts in smears of cattle feces, and Tzipori et al. (1980) used it to detect oocysts in human stools. Other Romanowsky stains such as Jenner’s stain have been used. The acid-fast Ziehl–Neelsen (ZN) stain used by Henriksen and Pohlenz (1981) to detect oocysts in calf feces was modified by Casemore et al. in 1984 (modified Ziehl–Neelsen, mZN) and became widely used to stain oocysts in fecal smears (Casemore, 1991). Many modifications to the mZN method have been published. The specific method often reflects operator preferences rather than differences in sensitivity and specificity. The fluorogenic stain, auramine phenol (AP), and negative staining with strong carbol fuchsin (Casemore et al., 1984, 1985; Casemore, 1991) or potassium permanganate (KMnO4; Fleck and Moody, 1988) have been widely used to stain oocysts in fecal smears. AP is more sensitive than mZN (Casemore et al., 1985) because it stains oocyst outer walls as well as internal structures. Phenol accelerates AP penetration through oocyst walls. In the author’s laboratory, AP and negative staining with KMnO4 is preferred. Oocysts and sporozoites can be readily seen with the fluorescein isothiocyanate (FITC) filter and ultraviolet (UV) filters of an epifluorescence microscope. Saffranin methylene blue stains oocysts a vivid orange-pink (Baxby et al., 1984). It is more sensitive than mZN and can be used to stain paraffin-embedded material. FITC-labeled monoclonal antibodies (mAbs) reactive with surface-exposed oocyst epitopes can also be used to detect oocysts in feces. Using AP and negative staining with strong carbol fuchsin, Cryptosporidium oocysts appear ringlike (4 to 6 µm in diameter) and exhibit a characteristic bright fluorescence against a dark red background. Using mZN, oocysts stain red and appear as spherules against a pale green background. The degree and proportion of staining varies with individual oocysts. Internal structures take up stain to varying degrees. Some appear amorphous, whereas others have the characteristic crescentic forms of sporozoites. Difficulties arise in discriminating between Cryptosporidium oocysts and other spherical objects of similar size, particularly with mZN, when only a few oocysts are present. Yeasts and fecal debris stain a dull red with mZN. Some bacterial spores might be acid-fast, but are too small to cause confusion. Cyclospora cayetanensis and Isospora belli oocysts also stain red, but are much larger. Storage of oocysts in formalin can compromise mZN staining. Using Jenner or Giemsa stains, oocysts appear semitranslucent and stain blue to azure. They might contain four to six red or purple dots. “Ghost” forms with a frosted-glass appearance and usually devoid of granules might also be found. A narrow, clear halo might surround the oocyst. For samples containing few oocysts, the use of an initial screening method such as AP followed by a confirmatory method such as immunofluorescence can augment confidence in diagnosis. Where fluorescence microscopy is not available, screening using mZN and confirmation using Giemsa or Jenner stains is useful but time consuming. Both morphology and specific staining characteristics of oocysts must be considered when attempting to make a laboratory diagnosis of cryptosporidiosis. Cryptosporidium-positive fecal samples should be available when personnel are familiarizing themselves with the staining techniques. When obtained from routine submissions, stool samples containing Cryptosporidium-oocysts can be stored at 4°C in either 2.5% potassium dichromate (K2Cr2O7) or 10% formalin for quality assurance and reference purposes.
A. 1.
Detection of Cryptosporidium Oocysts in Fecal Smears by Auramine Phenol Scope of Test
This rapid procedure is suitable for symptomatic cases. Oocysts are clearly visible against a dark background under the low-power objective (10 or 20×) of a fluorescence microscope. Oocysts should be confirmed under a higher-power (40, 50, or 100×) objective and measured under the 100× objective. Oocysts can be scraped from AP-stained microscope slides (Amar et al., 2001) for subsequent DNA extraction (Section XIIA1). The threshold of detection for this method is low (Section IX). Images of AP stained oocysts appear in Figure 6.1.
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187
Materials
Disposable gloves, glass microscope slides, Coplin jars or similar, methanol (95%), fluorescence microscope equipped with FITC (excitation 490 nm; emission 510 nm) and ultraviolet (excitation 355 nm; emission 450 nm) filters. Auramine phenol [Lempert] solution, 3% HCl in 70% methylated spirit, 0.1% potassium permanganate. Always include a positive control slide.
b.
Method 1. 2. 3. 4. 5. 6. 7. 8.
Fix the air-dried direct* or concentrate smear in methanol for 10 min. Immerse the slide in auramine phenol stain for 10 min. Rinse the slide in tap water to remove excess stain. Decolorize the slide in 3% HCl in methylated alcohol for 5 min. Rinse the slide in tap water. Immerse the slide in potassium permanganate for 30 s. Rinse the slide in tap water. Air-dry the slide and examine for the presence of oocysts using a fluorescence microscope equipped with FITC filters. 9. Measure the size and shape of the fluorescent bodies.**
2. a.
Preparation of Auramine Phenol (AP) Materials
Disposable gloves. For 1 L: 5-g Auramine O, 32-g phenol detached crystals (or 40 mL of 80% phenol); 320-mL industrial methylated spirits (IMS 90, or other alcohol); distilled water.
b.
Method 1. 2. 3. 4.
Dissolve phenol in methylated spirits (IMS 90, or other alcohol). Dissolve the Auramine O in the phenol–alcohol mixture. Make up to 1 L with distilled water. Store at room temperature in a lightproof glass bottle.
The protocol for AP and negative staining with strong carbol fuchsin differs slightly from the preceding protocol. The smear is fixed in methanol for 3 min, immersed in AP stain for 5 min, rinsed in tap water to remove excess stain, counterstained in cold strong carbol fuchsin for 10 s, and rinsed in tap water to remove excess stain prior to air drying.
B. 1.
Detection of Cryptosporidium Oocysts Using Modified Cold Strong Ziehl–Neelsen Stain Scope of Test
This procedure is suitable for symptomatic cases. Oocysts stain variably against a pale green background under the low-power objective (20×) of a bright field microscope. Oocysts should be confirmed under a higher-power (40, 50, or 100×) objective and measured under the 100× objective. Oocysts can be scraped from mZN-stained microscope slides (Amar et al., 2001) for subsequent DNA extraction (Section * The best smear thickness is achieved when either the hands of a watch or the print on a page can just be seen through the preparation. Smears should be air-dried avoiding the use of excessive heat, which can distort oocysts (see Section IV). ** Cryptosporidium oocysts appear ring-shaped (see Table 6.1 for oocyst dimensions) and exhibit a characteristically bright green fluorescence against a dark background.
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XII A1). The threshold of detection for this method is low (Section IX). Images of mZN stained oocysts appear in Figure 6.1.
a.
Materials
Disposable gloves, glass microscope slides, Coplin jars or similar, bright field microscope, immersion oil, methanol (95%), cold strong carbol fuchsin, 0.4% malachite green (w/v in distilled water), 1% HCl (v/v) in methanol (94%), and immersion oil. Always include a positive control slide.
b.
Method 1. 2. 3. 4. 5. 6. 7. 8.
Fix the air-dried direct* or concentrate smear in methanol for 3 min. Immerse the slide in cold strong carbol-fuchsin and stain for 15 min. Rinse the slide thoroughly in tap water. Decolorize the slide in 1% HCl (v/v) in methanol for 10 to 15 s. Rinse the slide in tap water. Counterstain the slide with 0.4% malachite green for 30 s. Rinse the slide in tap water. Air-dry the slide, and scan the slide using the 40× objective lens. Confirm the presence of oocysts under the oil immersion objective lens. 9. Measure the size and shape of the red-stained bodies.**
VIII. Immunological Methods A.
Antigen Detection Using Antibodies Labeled with Fluorescent Reporters
Direct and indirect fluorescent antibody detection using mAbs against Cryptosporidium oocyst wall antigens (C-mAbs) are specific and sensitive methods for detecting oocysts in fecal smears (Sterling and Arrowood, 1986; Stibbs and Ongerth, 1986; Garcia et al., 1987; McLauchlin et al., 1987; Arrowood and Sterling, 1989) and in environmental samples (Ongerth and Stibbs, 1987; Smith and Grimason, 2003). Many mAbs recognizing epitopes on the oocyst surface have been developed and commercialized. Direct and indirect fluorescent antibody detection can be more sensitive than conventional staining methods (Stibbs and Ongerth, 1986, Arrowood and Sterling, 1989). Commercially available FITC-C-mAbs are used routinely for detecting and enumerating oocysts in environmental samples (Smith and Rose, 1990; Smith and Grimason, 2003), and a variety of commercial kits are available for detecting oocysts in fecal and environmental samples (Table 6.2). When antibody paratopes in FITC-C-mAbs bind to epitopes on the oocyst surface, the visualized fluorescence defines the dimensions of the oocyst, enabling morphometric analyses. FITC-C-mAb can also bind to areas delineated by surface folds (where the oocyst wall has collapsed on itself), resulting in intense fluorescence at these sites. There is no commercially available antibody preparation specific for epitopes on human pathogenic or livestock pathogenic Cryptosporidium oocysts. Different mAbs used commercially recognize different sets of surface epitopes (Smith and Ronald, 2001); thus, the intensity of fluorescent emissions can be dependent on the oocyst isolate in question and the FITC-C-mAb present in the kit. Most, if not all, commercially available mAbs have been raised against epitopes on/in a limited number of C. parvum oocyst isolates (Table 6.3), and as the expression of exposed oocyst epitopes within the genus Cryptosporidium is expected to vary, it is likely that oocysts of different species and genotypes as well as some * The best smear thickness is achieved when either the hands of a watch or the print on a page can just be seen through the preparation. Smears should be air-dried avoiding the use of excessive heat, which can distort oocysts (see Section IV). ** Cryptosporidium oocysts stain red and appear as spherules (see Table 6.1 for oocyst dimensions) on a pale green background, but the degree and proportion of staining varies with individual oocysts.
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isolates within species and genotypes will fluoresce less intensely. This should be borne in mind when investigating clinical and environmental samples. Purified C. parvum and C. hominis oocyst isolates (n = 100) were found to stain variably from weak to strong with three commercially available FITC-CmAbs (unpublished). No correlation between fluorescence intensity and storage time was observed. Worryingly, these kits failed to detect 3% of Scottish C. parvum and C. hominis isolates, but the unstained isolates differed with the different kits. If this is suspected, a sensible approach would be to use diagnostic methods based on different principles (e.g., conventional staining, PCR-based methods). The three criteria recommended for the identification of Cryptosporidium oocysts by immunofluorescence are (1) characteristic apple-green fluorescence delineating the oocyst wall, under the FITC filter set; (2) round or slightly ovoid objects; and (3) a size of 4 to 6 µm in diameter, for most human pathogens (oocyst measurements in Table 6.1). Some kits include Evans Blue, which reduces nonspecific fluorescence and generates a red background fluorescence. Nonspecific fluorescence is yellow. Reference should always be made to the positive control to ensure that the size, shape, and color of putative oocysts are consistent with those of the positive control. A blue filter block (excitation, 490 nm; emission, 510 nm) is used to visualize FITC-C-mAb localization and ultra-violet (UV) excitation (excitation, 355 nm; emission, 450 nm) can be used to determine the presence of 4′,6-diamidino-2-phenyl indole (DAPI) stained sporozoite nuclei, if DAPI is used. Oocysts can be scraped from FITC-C-mAb (and DAPI) stained microscope slides (Nichols et al., 2006b) for subsequent DNA extraction (Section XIIA1). Images of FITC-C-mAb-stained oocysts appear in Figure 6.1.
B.
Antigen Detection Using Antibodies Labeled with Enzyme Reporters
Initially, enzyme immunoassay (EIA) and enzyme-linked immunosorbent assay (ELISA) systems differed in assay design, but both techniques are based on the principles of separating bound from unbound reagents, to increase assay specificity, sensitivity, and precision, and the detection of the analyte by antibody, using an enzyme as the reporter molecule. EIAs are useful because: (1) substances, such as antibodies or antigens, can be passively adsorbed to solid surfaces and (2) because one of the reactants is attached to the solid phase, the separation of bound and free reagents is easily undertaken. The major reasons why the ELISA format predominates in infectious diseases diagnostics include: (1) passive adsorption to plastics in the form of 96-well microtiter plates (or 8 well strips) enables easy manipulation and dispensing of reagents, the use of small volumes of reagents, and the potential for handling large numbers of samples rapidly, and (2) the color reaction can be read by eye, or automated using specially designed multichannel spectrophotometers, enabling electronic transfer of data that can be analyzed by statistical packages. Commercial kits based on enzyme-labeled antibodies to detect Cryptosporidium antigens are available (Table 6.2). EIA can offer increased specificity, sensitivity, and precision through the use of mAbs, and practicability, such as reagent stability, individual performance requirements, assay times, and automation potential.
1.
Enzyme Immunoassays
Cryptosporidium antigens can be sought in fecal samples with or without oocyst concentration. ELISA and immunochromatographic (IC) formats are the applications used most for commercial kits (Table 6.2). The sandwich ELISA is based on the principle that an antibody bound onto wells of 96 well microtiter plates or strips binds to and captures Cryptosporidium antigen in fecal suspensions. A second antibody binds to exposed epitopes on the captured antigen, sandwiching it between two antibodies. This reaction can be visualized in two ways: either the second antibody can be labeled with an enzyme reporter or a third antibody, which binds to the second antibody, is labeled with the enzyme reporter. The use of the third, enzyme-labeled antibody can increase the sensitivity of the assay. The reaction is visualized following the addition of a substrate that is catalyzed by the enzyme to produce a soluble color whose intensity can be assessed visually or by spectrophotometer. Some commercial ELISA kits are better at detecting infection than are some IF kits, although reports from different research groups vary (Graczyk and Cranfield, 1996; Garcia and Shimizu, 1997).
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Clinical samples can be dispatched to diagnostic laboratories either unfixed or fixed in 10% formalin, SAF, PVA, or other fixatives. The diagnostician should always determine whether any specific fixative interferes with the accuracy of the test method employed. These are normally listed in the inserts of commercial kits. The suitability of antigen detection methods for any given laboratory depends on the relative prevalence of Cryptosporidium infections, the number of specimens processed daily, and the balance between assay cost and time saved. Copro-antigen detection immunoassays are less time consuming and easier to perform than microscopic methods, and do not require experienced microscopists. As noted in Section VIIIA, not all commercial antibodies recognize all C. parvum and C. hominis oocyst antigens, and although both soluble and insoluble Cryptosporidium antigens are detected by EIA, the antibodies used in EIA kits may not react, or react weakly with the antigens of some Cryptosporidium species that are genetically distant from C. parvum and C. hominis. If this is suspected, a sensible approach would be to use diagnostic methods based on different principles (e.g., conventional staining, PCR-based methods). Copro-antigen detection immunoassays appear to offer no increase in sensitivity over microscopy. One advantage that copro-antigen detection immunoassays have over microscopy is that they can be invaluable in cases of infection in the absence of detectable oocysts (Section VA6). Specificity is reported to be high (98 to 100%).
2.
Immunochromatographic Assays
In the sandwich ELISA (VII B1), the speed of antigen–antibody interaction is determined by the molecular diffusion of the antigen and capture antibody, normally about an hour per reaction. In lateralflow immunochromatography, all fluids are drawn by wicking action through a membrane enclosed in a cassette. Soluble Cryptosporidium antigens in the test sample are drawn through the membrane, come into contact with, and bind to immobilized antibodies, which dramatically increases the speed of antigen–antibody interaction. Positive reactions are qualitative and are seen as a band of color at a specific location on the membrane, normally identified by a line on the cassette. Assay format can vary between kits. As for antigen detection by ELISA, the diagnostician should always determine whether any contraindications apply to the use of a commercial test and any fixative (e.g., PVA). Immunochromatographic assays provide diagnostic laboratories with a convenient alternative method for performing antigen detection assays for Cryptosporidium on stool samples. Specificity is reported to be high (98–100%). Some (e.g., Chan et al., 2000) report equal or better sensitivity than oocyststaining methods, whereas others (e.g., Johnston et al., 2003; Weitzel et al. 2006) report reduced sensitivity compared to conventional microscopic methods. Immunochromatographic assays can be invaluable in cases of infection in the absence of detectable oocysts (Section VA6).
IX. Sensitivity of Detection in Feces The sensitivity of detection is low using the methods previously described. For unconcentrated fecal smears, a detection limit of 106 oocysts per ml of feces was reported using Kinyon mZN (KmZN). After stool concentration, between 1 × 104 and 5 × 104 oocysts per g of unconcentrated stool are necessary to obtain a 100% detection efficiency using either KmZN or a commercially available FITC-C-mAb method (Weber et al., 1991). For bovine feces, the threshold for 100% detection efficiency using AP or FITC-C-mAb was 1 × 103 oocysts per g (Webster et al., 1996). Variations in fecal consistency influence the ease of detection, with oocysts being more easily detected in concentrates made from watery, diarrheal specimens than from formed stool specimens, because oocysts excreted in diarrheal stools are less likely to become attached to particulates than oocysts excreted in partially formed or formed stools and are more readily concentrated. The use of a FITC-C-mAb appears to offer no significant increase in sensitivity over conventional stains. In addition to microscopic techniques, antigen capture ELISAs have been reported with detection limits in the region of 3 × 105 to 106 oocysts per ml, which is similar in sensitivity to microscopy (Webster et al., 1996). Antigenic variability between clinical isolates of Cryptosporidium could compromise immunodiagnostic tests further. Although effective for diagnosing symptomatic infection, these methods are insensitive and unable to detect the smaller number of oocysts that might be
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TABLE 6.4 Advantages and Disadvantages of Diagnostic Methods for Identifying Cryptosporidium Oocysts Stain
Sensitivity
Specificity
Rapidity
Ease of Identification
Stool Consistency
Cost-Effectiveness
mZN AP IF EIA IC
+ ++ +++ +++ +++
+ ++ +++ +++ +++
++ +++ ++ +++ ++++
++ +++ +++ +++ ++++
Better with liquid stools Better with liquid stools Better with liquid stools Stool comminution required Stool comminution required
++ +++ + ++ ++
Note: mZN = modified Ziehl–Neelsen; AP = auramine phenol; IF = immunofluorescence; EIA = enzyme immunoassay; IC = immunochromatographic assay; + = moderate; ++ = good; +++ = very good; ++++ = excellent.
present in asymptomatic infections. In addition, anecdotal evidence suggests that smaller or minimal numbers of oocysts might be excreted in the chronic disease stages in immunocompromised individuals, indicating that oocyst detection methods might not be the ideal option for all situations. Table 6.4 summarizes some of the advantages and disadvantages of various fecal examination methods.
X.
Antibody Detection
Laboratory evidence of exposure/infection can be obtained indirectly following the analyses of various bodily fluids, such as serum/plasma, saliva, and feces (coproantibody) for the presence of antibodies to Cryptosporidium-specific antigens. In the absence of corroborative clinical or laboratory evidence, analysis for antibodies to Cryptosporidium-specific antigens provides evidence of previous or current exposure and/or indirect evidence of infection. Thus, the demonstration of specific circulating antibodies is only of diagnostic benefit if seroconversion from negative to positive serology or elevation in titer can be ascribed to a current or recent infection. In the absence of demonstrating seroconversion, titer elevation, or antibody isotype switching, positive reactions can indicate either current infection, past exposure, or both. Accordingly, serological methods tend to be used for epidemiological purposes. Most serological tests used for seroepidemiological surveys of exposure are ELISA or enzyme-linked immunoelectrotransfer blot (EITB; Western blot) based, using various aqueous extracts of C. parvum oocysts. Recently, ELISAs, using recombinant and partially purified C. parvum antigens, have been described. Following infection, a characteristic serum IgG antibody response to the 27-kDa and 15/17-kDa antigens can be demonstrated by EITB. Both the 27-kDa and 15/17-kDa antigens are associated with the sporozoite surface (Mead et al., 1988), and mAb neutralization of an epitope of the 27-kDa antigen reduced C. parvum infection in mice (Perryman et al., 1996). Further analysis of the 17-kDa and 27kDa antigens revealed that they are complex families of related proteins, some membrane associated and others soluble (Priest et al., 1999). Their surface association and the reduction in infection following neutralization of a 27-kDa antigen indicate that these antigens might be important targets for the protective humoral immune response. Therefore, both the 17-kDa and 27-kDa antigens appear to be functionally important candidate antigens for inclusion in seroepidemiological studies.
A.
Enzyme-Linked Immunoelectrotransfer Blot (EITB)
Changes in intensity of IgG and IgA responses to the 17/15-kDa antigen group, IgM responses to the 27kDa antigen group, and IgG responses to the 27-, 17/15-kDa antigen groups from C. parvum oocysts were detected between acute and convalescent sera from 10 confirmed cryptosporidiosis cases, based on (1) their reported symptoms, (2) oocyst-positive stools, and (3) EITB results (Moss et al., 1998). EITB appeared to be more predictive than ELISA. Sera from only 4 of the 10 cases demonstrated a significant change in IgG responses between acute and convalescent serum samples by ELISA (Moss et al., 1998). This led to further investigations of antibody isotype responses to these antigens to determine the benefits and usefulness of IgG antibody EITB for seroepidemiological surveys. The intensity of IgG antibody responses (which
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may approximate to a titer) to 27-kDa and 15/17-kDa antigens were determined, using computer-imaging densitometry software (Frost et al., 1998, 2003b), and antibody responses to the 27-kDa antigen, but not the 15/17-kDa antigen, declined over a 2-year time period (Frost et al., 1998). IgG antibody responses increased with age, and increased age appears to be predictive of a more intense response to each antigen group, probably reflecting increased exposure to either infection, oocyst/other parasite antigens, or both. Thus, the presence and intensity of IgG antibody responses to the 27-kDa and 15/17-kDa antigen groups predict a reduced risk of cryptosporidiosis (Frost et al., 2000a). The relationship between infection, determined by oocyst-positive stools, and IgG antibody positivity to the 27-kDa and 15/17-kDa antigens by EITB remains to be clarified fully. Not all symptomatic, infected, adult human volunteers, given doses of C. parvum oocysts ranging from 30 to 1 × 106, were IgG antibody positive to the 27-kDa and 15/17-kDa antigens by EITB (Moss et al., 1998), although 10 of 11 symptomatic volunteers were deemed blot-positive if IgM, IgG, or IgA antibody reactivity to either the 27-kDa or the 15/17-kDa antigens, or both, was demonstrated. Antibody responses to the 27-kDa and the 15/17-kDa antigens appear to characterize infection and may influence the development of signs and symptoms of disease (Moss et al., 1998). Uniform percentage large-format and minigel format can resolve the 27-kDa antigen, but gradient sodium dodecyl sulfate polyacrylamide (SDS-PAGE) gels resolve the 15- and 17-kDa bands better than uniform percentage SDS-PAGE gels and SDS-PAGE minigels. As both these 15- and 17-kDa bands appear to be diagnostic, the lower resolving power of minigels for these bands is not a drawback, and as they comigrate, they are referred to as the 15/17-kDa antigens, and antibody responses to them can be measured as a uniform reaction. Thus, both large-format and minigel analysis are similarly capable of detecting the 15/17kDa antigens, but for the 27-kDa antigen, the large-format gel detects more positive responses than the minigel (Frost et al., 2003b). However, given the appropriate quality control procedures, minigel assays generate findings comparable to those for the large-format gel assay at a fraction of the cost. As determining the intensity of the insoluble colorimetric response is paramount, care should be taken to minimize variability and to ensure that the antigen is uniformly applied across the blot, which results in the development of an even intensity of color across each band. Care should be taken to ensure that background staining is minimized on each blot and between blots. Computer imaging of blots should be performed shortly after the chromogen has completely developed, because insoluble color tends to fade over time. This avoids underestimating the coefficient of variation (Frost et al., 2003b). Comparison of antibody intensity to the 27-kDa and 15/17-kDa antigens has been used for retrospective and prospective epidemiological studies to monitor infection risk. Examples include the serological evidence of Cryptosporidium infections in southern Europe (Frost et al., 2000a), the serological analysis of a cryptosporidiosis epidemic (Frost et al., 2000b), serological evidence of endemic waterborne Cryptosporidium infections (Frost et al., 2000c, 2002b), and the serological responses to Cryptosporidium antigens among users of surface- versus groundwater sources (Frost et al., 2003a).
B.
Enzyme-Linked Immunosorbent Assay Using Recombinant (and Other) Proteins
Although ELISA, using crude C. parvum extracts, is less predictive than EITB (Moss et al., 1998), there has been renewed interest in ELISAs because the availability of recombinant, and other specific, Cryptosporidium antigens. A perceived drawback of EITB is the quantitation of the intensity of the IgG antibody response at both 27-kDa and 15/17-kDa bands. The ELISA format can circumvent these quantitation issues, as the resultant color change, caused by the converted substrates, remain soluble, so that spectrophotometric reading is optimized. Further benefits of ELISA include high sample throughput for population-based studies, reduced time and reagent costs, and its greater acceptance and use in diagnostic and reference laboratories, globally. A recombinant protein (rCP41, cloned and expressed in Escherichia coli), whose native homologue appears to be associated with the oocyst wall and whose gene sequence has been identified in the genomes of various Cryptosporidium species (Jenkins et al., 1999), is as effective as C. parvum crude oocyst antigen in an IgG antibody ELISA used to determine human seropositivity (Kjos et al., 2005). Further round robin testing of rCP41 should help determine its usefulness as a replacement for C. parvum crude oocyst antigen.
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The characteristic serum IgG antibody response to the immunodominant 27-kDa and 15/17-kDa antigens has been exploited to develop ELISA-based serodiagnostic and seroepidemiological assays. Results using a Cp23 (the recombinant form of the C. parvum 27-kDa antigen) IgG ELISA are comparable to analysis of IgG antibody reactivity to the 27-kDa antigen on EITB (Priest et al., 1999, 2001), and Cp23 ELISAs have been used to determine age-specific seroprevalence (Cox et al., 2005) and longitudinal infection trends (Ong et al., 2005) in humans as well as the presence of antibodies in serum and feces from neonatal calves (Wang et al., 2003). An IgG antibody ELISA using a partially purified (Triton X-114 phase-partitioned) form of the native 17-kDa protein from C. parvum oocysts as antigen (Priest et al., 1999, 2006) appears to be comparable to IgG antibody reactivity to the 15/17-kDa antigen on EITB. Results of the Cp23 ELISA and the enriched, native 17-kDa ELISAs can be used to determine age-specific seroprevalence and longitudinal infection trends. The magnitude of antibody responses is related to age and the number of previous infections. Importantly, although these antigens are derived from C. parvum, antibodies were detected from humans infected with C. parvum, C. felis, C. meleagridis, and with four different C. hominis subtype families (Priest et al., 2006). However, antibodies tested in immunoassays based on recognition of C. parvum-derived antigens may not react, or react weakly with these epitopes, particularly if the antibody response is to infecting species or genotypes genetically distant from C. parvum (see Sections VIIIA and VIIIB1). Results obtained by EITB and both the Cp23 and the enriched, native 17-kDa ELISAs indicate that these ELISAs show promise. However, their usefulness as an adjunct to routine diagnosis must await testing of further diverse samples and the standardization of antigens.
XI. Biopsy Before 1980, human cryptosporidiosis was diagnosed histologically by finding life-cycle stages in the microvillus region of the intestinal mucosa obtained by biopsy or at necropsy. In haematoxylin- and eosin-stained sections, these stages appeared as small, spherical bodies (2 to 5 μm) in the microvillus region (Figure 6.2b). Special staining procedures offered little diagnostic advantage over haematoxylin and eosin. Transmission electron microscopy was used for confirmation and revealed distinct features of life-cycle forms (Chapter 1). Neither light microscopy nor TEM examination of tissue are now used for routine diagnosis.
FIGURE 6.2 Images of sections of uninfected (Image A) and infected (Image B) neonatal mouse intestines. In image B, the mouse was infected with 30 C. parvum oocysts. Arrows point to the life-cycle stages localized intracellularly, yet extracytoplasmically, within the brush border.
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Biopsy specimens require invasive procedures and careful sample processing. Not all regions of the intestine are infected, which can give rise to sampling errors. This invasive, expensive, and timeconsuming technique has been replaced with alternative techniques. Biopsy and light or electron microscopy are still useful when investigating histopathological and cytoarchitectural changes associated with infection.
XII. Molecular Diagnosis—Nucleic Acid Detection Methods PCR-based methods are more sensitive than conventional and immunological assays for detecting oocysts in feces, although the sensitivity of published methods can range from 1 to 106 oocysts. Identifying the species infecting humans and animals is important in determining the epidemiology of disease and likely transmission routes. Molecular methods are often restricted to specialist laboratories, but are necessary to determine Cryptosporidium species/genotypes and subtypes.
A.
Extraction of Cryptosporidium DNA from Oocysts in Clinical and Environmental Samples
Stools, sputum, bile, mucoid secretions, tissue biopsies, and paraffin-embedded sections can be submitted to clinical laboratories for species and subspecies identification, but the majority of samples are stools. On occasion, mZN, AP, or IFA stained and unstained smears on microscope slides will be submitted. No recommended method for extracting DNA from oocysts exists, and the sensitivity of most methods described has not been addressed fully. Table 6.5 identifies methods used for extracting oocyst DNA from feces where seeding and/or sensitivity data are available. DNA can be extracted either after partial purification of oocysts using flotation, sedimentation, or IMS techniques, or from oocysts in feces following zirconia bead extraction (McLauchlin et al., 1999). If concentration by formol ether sedimentation is the routine laboratory procedure, oocyst concentrates suitable for lysis and amplification by PCR can be made by substituting grade 1 water for the 10% formalin normally used (Nichols et al., 2006a). DNA loss can be a consequence of subsequent DNA purification using commercial purification columns, but cleaner products are generally generated and normally there is an adequate number of oocysts present in the sample to extract sufficient Cryptosporidium DNA for PCR-RFLP and sequence analysis.
1.
DNA Extraction for Outbreak and Epidemiological Investigations
In some instances, rapid oocyst isolation and extraction methods are required for outbreak investigations, particularly when nested PCR-based methods are used. A small-scale formol/water ether method described in Section VIA was developed for outbreak investigations, particularly when numerous molecular analyses were required (Nichols et al., 2006b). Similarly, oocysts can also be scraped from air-dried unfixed, unstained smears and mZN-, AP-, and IF- (Amar et al., 2001; Nichols et al., 2006) stained microscope slides, responsible for the original diagnosis, for subsequent DNA extraction. The slidescraping method of Amar et al. (2001) used fecal samples stored at 4°C for up to 2 years without preservatives. Methanol fixed smears were stained with mZN, AP, or IF and 2 weeks later, immersed in a guanidinium thiocyanate-based lysis buffer, the oocysts were removed by rubbing the stained smear vigorously with a cotton swab. DNA extraction and PCR are described in Table 6.4 (McLauchlin et al., 1999). Similarly, IMS can be used to concentrate oocysts from aqueous extracts of stools for subsequent DNA extraction, particularly when oocyst abundance is low. Protocols for extracting DNA from histological slides usually follow conventional procedures. Prior to routine adoption in clinical laboratories, both the variability between methods and the recognized difficulties in amplifying nucleic acids from fecal specimens by PCR must be overcome. Fecal samples can contain many PCR inhibitors. In addition to bilirubin and bile salts, complex polysaccharides
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TABLE 6.5 Some Methods for Extracting Cryptosporidium Oocyst DNA from Feces PCR
Matrix
DNA Extraction and Purification Methods from Oocysts
Facilitator
QIAamp® DNA stool Inhibit-EX-tablet minikita, boil 5 min, freeze-thaw 3× (liquid nitrogen for 1 min, boil for 2 min) Frozen pig and Freeze-thaw 3× (liquid Bovine serum calf stools nitrogen for 2 min, 75°C albumin (BSA) (diluted 1:4 w/v for 2 min), centrifuge, in PBS) and add lysis buffer (Tris-EDTA-SDSproteinase K) to pellet, glassmilkb Human stool Modified FastDNA Prep PVP in extraction (0.3–0.5 g) kitc and QIAquick spin method. 11.3% preserved in column (24 of 213 2.5% potassium samples) were dichromate inhibitory Unpreserved Water-ether extraction BSA, Tween 20 solid (0.4–0.5 g) and washes in lysis and PVP or liquid (200 buffer (Tris-EDTAincorporated into µL) human SDS),d freeze-thawing PCR stool 15× (liquid nitrogen for 1 min, 65°C for 1 min) 200 µL of whole Oocyst disruption by PVP in extraction human stool shaking in guanidinium method. thiocyanide and zirconia beads with the addition of isoamyl alcohol, to prevent foaming. DNA extraction and purification following (Boom et al., 1990). Unpreserved human stool (180–200 mg)
a b
c
d
Target (Detection Limit) Nested COWP (5 × 102 oocysts)
Sensitivity [(% positives); Number of Positives/Total] (97%) 86/89
Reference Bialek et al., 2002
Nested Laxer et al. Not available (1991) using 100 seeded oocysts
Ward and Wang, 2001
Direct 18S rRNA
(100%) 53/53 plus 31 negatives by microscopy
da Silva et al., 1999
Single-tube nested COWP and nested 18S rRNA
(97.8%) 90/92
Nichols et al., 2006a
Direct 18S rRNA and Direct COWP and Direct TRAP-C1
(97%) 204/218
(98.9%) 91/92
McLauchlin et al., 1999
(91%) 191/218 (66%) 139/218
QIAamp® DNA stool minikit is based on the use of proprietary buffers and DNA purification through a silica column. Glassmilk (GENECLEAN® Bio 101) is based on DNA adsorption to silica suspensions that allows subsequent washing of bound DNA to remove inhibitors. Modified FastDNA kit uses FP120 FastPrep Cell Disruptor to disrupt oocysts in a proprietary buffer that minimizes adsorption of DNA to fecal particles (Cell lysis/DNA Solubilizing Solution). Polyvynylpyrrolidone (final concentration of 0.5% w/v) used in this step precipitates polyphenolic compounds and the solubilized DNA is bound to the binding matrix in the presence of chaotropic salt, which is washed and then eluted. The final purification of DNA is performed in a QIAquick spin column. This method uses semipurified oocysts and no DNA purification. The presence of 0.5% SDS in the oocyst lysate is inactivated by the addition of 2% Tween 20 to the PCR mixture.
are significant inhibitors. Boiling fecal samples in 10% polyvinylpolypyrrolidone (PVP) before extraction and the use of 400 ng/µL of nonacetylated bovine serum albumin (BSA) in PCR can reduce inhibition.
2.
DNA Extraction from Small Numbers of Partially Purified Oocysts
Fifteen cycles of freeze-thawing was found effective in extracting DNA from small numbers (~10+) of partially purified oocysts (Nichols et al., 2003, 2006a, 2006b; Nichols and Smith, 2004). Oocysts of the
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C. parvum Iowa isolate, used by many researchers to develop molecular methods or for positive controls, were found very susceptible to freeze–thawing (Nichols and Smith, 2004). Oocysts of other species/ genotypes present in feces or environmental samples can be more resistant to freeze–thawing.
a.
Materials
Cold-resistant (for cryogenic temperatures) and disposable gloves, 1.5-mL microcentrifuge tubes, tube holder suitable for 1.5-mL microcentrifuge tubes,* cryogenic reservoir containing liquid nitrogen, water baths set at 65 and 55°C, oven set at 90°C, microcentrifuge (16,000 × g), lysis buffer (50-mM Tris/HCl, pH 8.5, 1-mM ethylene diamine tetra-acetic acid, pH 8, 0.5% sodium dodecyl sulfate** (SDS), proteinase K (at a final concentration of 200 μg per mL), and ice. Always include a positive control.
b.
Method
1. Suspend the partially purified oocysts (maximum volume = 90 µL) in 10 µL of 10× lysis buffer in a microcentrifuge tube. 2. Cap and insert each tube fully into the holes of a rack insert* or other holding device. 3. Gently lower tubes onto the surface of the liquid nitrogen.**** 4. Let them float on the liquid nitrogen for 1 min. 5. Gently raise the tubes out of the liquid nitrogen and float them immediately on the water in the 65ºC water bath. Leave for 1 min. 6. Repeat this process 14 times. Make a note of each cycle of freezing and thawing. 7. Add proteinase K (at a final concentration of 200 μg per mL) to each sample, recap each tube, and incubate for 3 h in a 55°C water bath. 8. Transfer microcentrifuge tubes to an oven set at 90°C for 20 min. 9. Chill microcentrifuge tubes on ice for 1 min. 10. Centrifuge microcentrifuge tubes at 16,000 × g for 5 min. 11. Uncap tubes and remove ~70 µL of supernatant.***** 12. Store each extract in a clean, capped, and labeled tube at 20°C until used for PCR amplification. Reagents for PCR reactions are dispensed in 0.5-mL thin-walled tubes. PCR reactions are set up in a designated laboratory in a hood presterilized by UV light. Each reaction is performed in either 50- or 100-μL containing premixed reagents at final concentrations of 200 μM of each of the four dNTP’s; BSA at 400 ng/µL; MgCl2 at concentrations varying from 2.5 mM; 2.5U of Taq polymerase; and Tween 20 at 2% and primers (0.2 µM) in 1× PCR buffer IV. About 2 or 3 μL of DNA template (for the primary PCR of the two-step nested PCR assay) of lysate (defrosted at room temperature, mixed by vortexing for 10 s, and centrifuged at 14,000 × g for 10 s in a microcentrifuge) are used for amplification. Secondary PCRs are set up by transferring 2 µL of primary PCR product to 100-µL total reaction volume following published protocols. Reagents and conditions for the primary and secondary PCRs are the same. Three negative controls are set up for each PCR run: one using the water for preparing the megamix (performed in the laboratory designated for pre-PCR manipulations), one using LB set up before dispensing the test samples, and one set up after all the test samples for an individual PCR run are dispensed. One positive * Tubes containing the samples are pushed fully into a tube rack or holder, ensuring that the meniscus of each sample protrudes below the base of the insert. The insert should be of a smaller diameter than the neck of the liquid nitrogen reservoir so that it can be floated on the surface of the liquid nitrogen (and water), thus ensuring that the samples become immersed in each fluid. The expanded polystyrene tube rack insert can be readily cut to shape to fit into the aperture of the cryogenic liquid nitrogen reservoir. ** SDS is inhibitory to Taq polymerase at concentrations as low as 0.01%; therefore, it is necessary to neutralize the SDS present in the extracted DNA, before PCR. The addition of 2% Tween 20 will neutralize up to 0.05% SDS. *** Observe local safety codes of practice when using liquid nitrogen. **** A pair of long forceps is suitable for transferring samples between each fluid. *****For important samples, a larger volume can be withdrawn as long as the pellet remains stable after centrifugation.
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control (preferably from a Cryptosporidium sp. unexpected in samples under analysis) of a known DNA concentration, which is appropriate to each PCR run, is set up as the last sample. PCR amplifications are performed following published amplification protocols. Secondary PCR product is visualized by gel electrophoresis either on 2 or 1.4% agarose gels, stained with ethidium bromide. Band intensities are classified according to the concentration of amplicons as negative PCR, faint band denoting positive PCR but insufficient amplicon concentration for RFLP analysis, 1+ denoting low-intensity bands with sufficient amplicon concentration for RFLP analysis, 2+ denoting medium-intensity bands with sufficient amplicon concentration for RFLP analysis, 3+ denoting strong-intensity bands with sufficient amplicon concentration for RFLP analysis, and 4+ denoting verystrong-intensity bands with sufficient amplicon concentration for RFLP analysis. The primers and step cycle protocols for amplifying either 18S rRNA gene fragments (Johnson et al., 1995; Nichols et al., 2003; Xiao et al., 1999, 2001) or the COWP gene fragment (Homan et al., 1999) are given in Table 6.6.
B.
Primer, Gene Locus Selection, and PCR
Care is necessary when choosing primers, because some primers amplify genus-specific DNA, whereas others amplify species-specific DNA. C. parvum and C. hominis are the most common species infecting humans, and mixed species infections are reported far less frequently. In livestock, the pathogens vary. A thorough understanding is required of the usefulness, strengths, and weaknesses of PCR-based amplification at each locus. Few studies have compared the efficacy of amplification and differentiation of species present. Jiang and Xiao (2003) evaluated the performance of 10 commonly used loci to detect and differentiate human pathogenic species of Cryptosporidium. The PCR-RFLP of all 18s rRNA gene loci tested amplified and differentiated all species tested (C. hominis, C. parvum, C. meleagridis, C. felis, C. canis, C. muris, and C. suis), but PCR-RFLP of the TRAP C1, COWP, HSP 70, and DHFR gene loci only amplified and differentiated C. hominis, C. parvum, and C. meleagridis. This might be sufficient for species present in the majority of human, but not animal, samples. The most robust information regarding species/genotype information has been derived from the study of three genetic loci (two 18S rRNA [Xiao et al., 1999, 2001; Johnson et al., 1995; Nichols et al., 2003] and the Cryptosporidium Oocyst Wall Protein [COWP; Spano et al., 1997; Homan et al., 1999]) gene fragments by PCR-RFLP and/or sequencing amplicons. Methods targeting 18S rRNA gene loci have been used extensively and globally for the identification of Cryptosporidium species in human and nonhuman hosts and environmental (water and food) samples. These methods are advocated here. A multilocus approach to characterizing Cryptosporidium isolates is essential for accuracy and increased confidence in diagnosis, and various 18S and other loci are available for species (and subtype) determination (see Chapter 5). Multiplexing, real-time PCR, and melting curve analysis offer prospects for multiple species and genotype detection in automated procedures. Subtyping tools including glycoprotein (GP) 60 gene sequencing, mini- and microsatellite markers, and analysis of a double-stranded RNA element can subtype C. parvum and C. hominis (Cacciò et al., 2005); however, the public/veterinary health value of subtyping tools depends on their ability to map transmission patterns in defined endemic foci and, as such, will require further research and consolidation.
1.
18S rRNA Gene Fragments
There is no “standard” genetic locus recommended for species identity, but RFLP or sequencing of 18S gene loci provide information about more species than the COWP gene locus. The nested assay (Xiao et al., 1999, 2001) is able to detect most Cryptosporidium species and genotypes by PCR-RFLP, and has been tested and validated in numerous laboratories. The external primers amplify a product of about 1325 bp, and the internal primers amplify a product of about 826 bp. PCR amplification of Cryptosporidium DNA using the 18S rRNA primers (CPB-DIAGF/R) of Johnson et al. (1995) yields products that vary in length from 428 to 455 bp. The Johnson et al. primers are included because they were evaluated for cross-reactions against a total of 23 microorganisms, and the primers have been
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TABLE 6.6 Step Cycle PCR Protocols for 18S rRNAa and Single-Tube Nested COWPb PCR Primers
Step Cycle Protocol
Reference
CPB-DIAGF AAG-CTC-GTA-GTT-GGA-TTT-CTG CPB-DIAGR TAA-GGT-GCT-GAA-GGA-GTA-AGG
80°C, 5 min; 98°C, 30 s; 1 cycle Johnson et al., 55°C, 30 s; 72°C, 1 min; 94°C, 30 s; 39 cycles 1995 72°C, 10 min; 1 cycle 4°C, soak
N-DIAGF2 CAA TTG GAG GGC AAG TCT GGT GCC AGC
N-DIAGF2/R2 (Primary PCR of two-step nested)
N-DIAGR2 CCT TCC TAT GTC TGG ACC TGG TGA GT
94°C, 5 min; 94°C, 1 min; 68°C, 30 s; 72°C, 30 s; 35 cycles 72°C, 10 min; 1 cycle 4°C, soak
CPB-DIAGF AAG-CTC-GTA-GTT-GGA-TTT-CTG CPB-DIAGR TAA-GGT-GCT-GAA-GGA-GTA-AGG
Nichols et al., 2003
CPB-DIAGF/R (Secondary PCR of two-step nested) 94°C, 1 min; 94°C, 1 min; 60°C, 30 s; 72°C, 30 s; 35 cycles 72°C, 10 min; 1 cycle 4°C, soak
XF1 (outer) TTC-TAG-AGC-TAA-TAC-ATG-CG XR1 (outer) CCC-ATT-TCC-TTC-GAA-ACA-GGA XF2 (inner) GGA-AGG-GTT-GTA-TTT-ATT-AGA-TAA-AG XR2 (inner) AAG-GAG-TAA-GGA-ACA-ACC-TCC-A
94°C, 3 min; 1 cycle Xiao et al., 1999, 94°C, 35 s; 55°C, 45 s; 72°C, 1 min; 35 cycles 2001 72°C, 7 min; 1 cycle 4°C, soak
oocry3 (outer) AGA-TTA-ACA-GAA-TGC-CCA-CCA-GGT-A oocry4 (outer) CCA-TGA-TGA-TGT-CCT-GGA-TTT-TGT-A oocry1 (inner) CCT-GGA-TAT-CTC-GAC-AAT Oocry2 (inner) GCG-AAC-TAA-TCG-ATC-TCT-CT
94°C, 5 min; 1 cycle 94°C, 1 min; 67°C, 1 min; 72°C, 1 min; 20 cycles 94°C, 1 min; 54°C, 1 min; 72°C, 1 min; 35 cycles 72°C, 10 min; 1 cycle 4°C, soak
a
b
Homan et al., 1999
From Johnson, D.W., Pieniazek, N.J., Griffin, D.W., Misener, L., and Rose, J.B. 1995. Development of a PCR protocol for sensitive detection of Cryptosporidium in water samples. Appl. Environ. Microbiol. 61, 3849–3855; Nichols R.A.B., Campbell B.M., and Smith H.V. 2003. Identification of Cryptosporidium spp. oocysts in UK still natural mineral waters and drinking waters using a modified nested PCR-RFLP assay. Appl. Environ. Microbiol. 69, 4183–4189. Xiao, L., Morgan, U., Limor, J., Escalante, A., Arrowood, M., Shulaw, W., Thompson, R.C.A., Fayer, R., and Lal, A.A. 1999. Genetic diversity within Cryptosporidium parvum and related Cryptosporidium species. Appl. Environ. Microbiol. 65, 3386–3391; Xiao, L., Sing, A., Limor, J., Graczyk, T.K., Gradus, S., and Lal, A.A. 2001. Molecular characterisation of Cryptosporidium oocysts in samples of raw surface water and wastewater. Appl. Environ. Microbiol. 67, 1091–1101. From Homan, W., Van Gorkom, T., Kan, Y.Y., and Hepener, J. 1999. Characterization of Cryptosporidium parvum in human and animal feces by single-tube nested polymerase chain reaction and restriction analysis. Parasitol. Res. 85, 707–712.
shown to work in a variety of matrices in many different regions of the world. The nested Nichols–Johnson 18S rRNA assay has also been shown to be sensitive (Nichols et al., 2003). For detecting small numbers of oocysts (<100) consistently, a nested PCR is required. Two 18S rRNA PCRs have been tested extensively in the author’s laboratory (Xiao et al., 1999, 2001; Nichols et al., 2003) and the Nichols–Johnson primers appear to be more sensitive, particularly with small numbers (≤10) of C. parvum, C. hominis, C. felis, and C. muris oocysts, probably because of the smaller amplicon size (Nichols et al., 2006a). Although less sensitive, the Xiao et al. (1999, 2001) assay has the benefit
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199 TABLE 6.7 Cryptosporidium spp. and Genotypes Determined by RFLP of the Amplicons Defined by the XR2/XF2 Primers following Digestion with Enzymes AseI (VspI) and Ssp1 Cryptosporidium Species
VspI
C. hominis C. parvum gene type A C. parvum gene type B C. muris C. andersoni C. felis C. canis C. suis C. wrairi C. baileyi C. meleagridis C. serpentis C. varanii C. hominis monkey Cryptosporidium mouse Cryptosporidium ferret Cryptosporidium kangaroo, koala
561, 628, 625, 731, 731, 476, 633, 632, 628, 620, 456, 729, 628, 559, 457, 457, 631,
104, 104, 104, 102 102 182, 102, 104, 104, 104, 171, 102 104, 104, 175, 174, 104,
SspI
102, 70 102 102
104, 102 94 102 102 102 104, 102 102 102, 70 104, 102 104, 102 102
449, 449, 449, 448, 448, 426, 417, 453, 449, 572, 449, 414, 418, 461, 449, 449, 441,
254, 254, 254, 385 385 390, 254, 365, 254, 254 254, 370, 255, 254, 254, 254, 254,
111, 12, 11 108, 12, 11 119, 9
33, 15 105, 33, 20 11, 9 109, 11, 11 108, 11, 33, 14 109, 33, 109, 11 112, 12, 111, 12, 109, 33
11 19 11 11
Source: From Xiao, L., Morgan, U., Limor, J., Escalante, A., Arrowood, M., Shulaw, W., Thompson, R.C.A., Fayer, R., and Lal, A.A. 1999. Genetic diversity within Cryptosporidium parvum and related Cryptosporidium species. Appl. Environ. Microbiol. 65, 3386–3391.
of being able to detect more species than the Nichols–Johnson assay by RFLP, particularly the important human pathogens C. parvum and C. hominis. Table 6.7 contains the Cryptosporidium spp. and genotypes determined by RFLP of the amplicon defined by the XR2 / XF2 primers following digestion with enzymes AseI (VspI) and SspI (Xiao et al., 1999). Table 6.8 contains the Cryptosporidium spp. and genotypes determined by RFLP of the amplicon defined by the CPB-DIAGR/F primers following simultaneous digestion with restriction enzymes VspI/DraI (Nichols et al., 2003), which can identify five major species. Table 6.9 contains the RFLP result of the same amplicons separately digested with the restriction enzymes VspI/AseI, SspI and DdeI according to available GenBank sequences of the 18S rRNA gene (Nichols et al., 2003). TABLE 6.8 Structural Analysis of the 18S rRNA Gene Defined by CPB-DIAGR Primers After Simultaneous Digestion with the Restriction Enzymes VspI or AseI and DraI Cryptosporidium Species (amplicon length in bp) C. C. C. C. C. C.
hominis (438) parvum (435) muris (431 or 432) felis (455) baileyi (428) meleagridis (434)
Number of VspI/AseI Sites ↓TAAT) (AT↓
Number of DraI Sites ↓AAA) (TTT↓
2 2 1 2 2 3
None None None 1 1 None
Fragments Length (bp) 222; 219; 319; 189; 128; 171;
112; 112; 112 112; 112; 112;
104 104 104; 50 104; 84 104; 47
Source: From Nichols R.A.B., Campbell B.M., and Smith H.V. 2003. Identification of Cryptosporidium spp. oocysts in UK still natural mineral waters and drinking waters using a modified nested PCR-RFLP assay. Appl. Environ. Microbiol. 69, 4183–4189. With permission.
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TABLE 6.9 Cryptosporidium spp. and Genotypes Determined by RFLP of the Amplicons Defined by the CPBDIAG Primers Following Digestion with Enzymes VspI or AseI, SspI and DdeI, According to Available GenBank Complete/Partial Sequences of the 18S rRNA Gene Cryptosporidium Species (amplicon length in bp) C. hominis (438) C. parvum (435) C. muris (432) C. andersoni (431) C. felis (455) C. canis (429) C. wrairi (435) C. suis (435) C. baileyi (428) C. meleagridis (434) C. serpentis (430) C. varanii (432) C. hominis monkey (436) Cryptosporidium mouse (439) Cryptosporidium ferret (438) Cryptosporidium koala (436) Cryptosporidium kangaroo (436)
VspI/AseI 222, 219, 320, 319, 239, 213, 219, 219, 212, 171, 318, 216, 220, 175, 174, 220, 220,
104, 104, 112 112 104, 104, 104, 104, 104, 104, 112 108, 104, 104, 103, 104, 104,
112 112
112 112 112 112 112 112, 47 112 112 112, 48 113, 48 112 112
SspI 264, 264, 395, 394, 401, 264, 264, 375, 264, 264, 380, 264, 264, 264, 264, 264, 373,
111, 40, 12, 108, 40, 12, 37 37 40, 14 105, 40, 20 109, 40, 11, 40, 11, 9 164 119, 40, 11 36, 14 109, 40, 19 109, 52, 11 112, 40, 12, 111, 40, 23 109, 63 63
GenBank Accession No.
DdeI 11 11
11
11
204, 201, 224, 265, 221, 195, 201, 201, 262, 200, 264, 198, 202, 205, 204, 202, 202,
166, 166, 166, 166 166, 166, 166, 166, 166 166, 166 166, 166, 166, 166, 166, 166,
68 68 42 68 68 68 68 68 68 68 68 68 68 68
L16997 L16996 AF093498 L19069 AF087577 AF112576 AF115378 AF108861 L19068 AF112574 AF093502 AF112573 AF112569 AF108863 AF112572 AF108860 AF112570
Not all Cryptosporidium species/genotypes can be differentiated by PCR-RFLP of the 18S rRNA loci. However, most of the species that are currently known to be human pathogens and commercially important for livestock can be identified by PCR-RFLP. For bovine samples, sequencing has been used to distinguish C. parvum from C. bovis or the Cryptosporidium deer-like genotype (Fayer et al., 2005), but they can also be readily differentiated by digestion with MboII (Feng et al., 2007). Sequencing of an amplicon can offer better information than PCR-RFLP, but sequencing is more expensive and takes longer than PCR-RFLP. Currently, sequencing availability varies in different parts of the world and, for the more common species that infect humans and livestock, PCR-RFLP has an important role to play.
2.
Cryptosporidium Oocyst Wall Protein (COWP) Gene Fragment
A single-tube nested PCR-RFLP assay (Homan et al., 1999) that amplifies a fragment of the gene coding for COWP distinguishes C. hominis from C. parvum and is recommended over that of Spano et al. (1997) for human samples. It is more sensitive and offers a solution to contamination frequently experienced in nested PCRs because of reamplification of PCR products. In this single-tube nested PCR, the inner and outer primers are added to the initial reaction mixture. Optimization of primer set concentrations and annealing temperatures result in the preferential amplification of one product size only, defined by the inner primers.
3.
MAS-PCR
A multiplex allele-specific PCR (MAS-PCR) based on the dihydrofolate reductase gene sequence differentiates C. hominis (357 bp) from C. parvum (190 bp) in a one-step reaction that can be distinguished on agarose gel (Giles et al., 2002). MAS-PCR is as sensitive as other assays targeting the 18S rRNA gene for C. hominis and C. parvum detection, and can detect more mixed (C. hominis and C. parvum) species infections than other PCR assays (Giles et al., 2002). The one-step MAS-PCR assay is simpler and less expensive to perform than RFLP-based methods because differentiation between C. parvum and C. hominis DNA relies on amplicon size without the requirement for endonuclease digestion and RFLP analysis, thus reducing assay time considerably. Of interest is the fact that MAS-PCR detected more mixed species infections than the other 18S rRNA gene assays tested. Based on assay sensitivity and time savings, MAS-PCR could be a valuable addition to the molecular tools used for human outbreak
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and large-scale epidemiological investigations. Furthermore, if MAS-PCR does detect more mixed C. parvum and C. hominis infections, it could modify our understanding of the epidemiology of human disease. However, MAS-PCR failed to detect DNA of C. felis, C. canis, C. muris, and C. suis in human samples (Jiang and Xiao, 2003), and its usefulness for determining the range of pathogenic species found in animal samples is probably limited. Therefore, for any molecular species-typing assay, it remains fundamental to determine the questions that need to be addressed before deciding which assays to use.
4.
Mixed Infections
Mixed infections of C. parvum and C. hominis appear to be uncommon, but have been widely reported (e.g., McLauchlin et al., 2000; Giles et al., 2002; Fretz et al., 2003; Tanriverdi et al., 2003; Mallon et al., 2003; Jiang and Xiao, 2003; Cama et al., 2006; Morse et al., 2007) using different molecular typing tools. PCR–RFLP of the 18S rRNA is used commonly for genotyping Cryptosporidium spp. because of its high sensitivity and ability to detect more species than PCR-RFLP tools based on other genes (Jiang and Xiao, 2003). The use of genus-specific molecular typing tools may not detect the presence of mixed Cryptosporidium species as readily as other typing tools because of preferential PCR amplification of the predominant genotypes or the specificity of the molecular tools chosen. One study demonstrated that an 18S rRNA PCR-RFLP tool had only a 31 to 74% success rate in detecting mixtures of C. parvum and C. hominis oocysts (Reed et al., 2002). Mini- and microsatellites used to differentiate C. parvum and C. hominis found that 12% of 135 clinical specimens from Aberdeenshire, Scotland, had mixed C. parvum and C. hominis infections (Mallon et al., 2003). Of 975 Scottish samples from the same time period, including those analyzed by Mallon et al., only 0.6% of samples contained mixtures, using the COWP species-typing tool. In epidemiological investigations, this may be further complicated by the analysis of a single sample, particularly if infection and oocyst excretion dynamics, clinical signs and symptoms, and health sequelae differ for different infecting species or genotypes. Furthermore, the inability to detect mixed species can compromise our understanding of the epidemiology of human and livestock disease and the health sequelae of cryptosporidiosis. In endemic areas of disease and in the immunocompromised, mixed infections may be more common than presumed. Mixed infections can be a consequence of sporadic infection, or of waterborne (McLauchlin et al., 2000) or foodborne outbreaks of disease if more than one infectious species or genotype is present in the contaminating vehicles. 18S rRNA-based PCR-RFLP tools have been widely accepted because they are sensitive and can detect and differentiate between a wide range of Cryptosporidium species and genotypes. However, Cama et al. (2006) found that their usefulness in detecting mixed infections was compromised by preferential PCR amplification of the dominant species or genotype in the specimens they tested. Preferential amplification of the dominant species or genotype is likely to occur with PCR; therefore, PCR tools with broad specificity in combination with species-specific tools is a better option to determine the frequency and duration of infection and the clinical and epidemiological relevance, particularly mixed infections of C. hominis and C. parvum in immunocompetent and immunocompromised hosts, in both endemic and nonendemic settings. Here, quantitative real-time PCR, using species-specific probes can offer significant advantages (Limor et al., 2002).
5.
Quantitative Real-Time PCR
Quantitative real-time PCR methods use patented technologies (e.g., TaqMan™, Applied Biosystems; LightCycler™, Roche Molecular Biochemicals). For both technologies, fluorogenic dyes are incorporated into the amplicon during PCR; thus, amplicon fluorescence increases as more PCR product is generated. Both systems can detect PCR products during the initial cycles of the PCR reaction when amplification is exponential, thus enabling quantitative analysis of fluorescent product. Quantitative real-time PCR methods have been used to identify different Cryptosporidium species by exploiting the genetic polymorphism of the 18S rRNA gene for devising probes with differing melting temperatures, and for the quantitative detection of Cryptosporidium oocysts in environmental water samples and sewage. The increased sensitivity of real-time PCR guarantees increased speed of detection and qualitative diagnosis;
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also, the quantitative nature of the assay will be invaluable in estimating levels of contamination. The “closed-tube” assay format reduces the danger of contamination from “carry over.” Quantitative realtime PCR should be a useful tool for the future, once problems related to matrix inhibition are overcome. Currently, there are no standardized quantitative real-time PCR methods.
6.
Typing and Subtyping for Disease and Source Tracking
The value of characterizing the genetic diversity of Cryptosporidium at different levels of specificity and the importance of appropriate nucleic acid analysis cannot be overemphasized. Molecular tools for inter- and intraspecies discrimination differ, and the publication of complete genomic sequences of C. parvum and C. hominis has assisted in developing appropriate typing and subtyping tools (Cacciò et al., 2005). Genetic markers differ in their information content, and the nature of the DNA fragment selected for detecting and characterizing Cryptosporidium should be considered carefully (see Table 6.10 below). To detect species, analysis of highly or moderately conserved coding regions (e.g., 18S ribosomal DNA, structural and house-keeping genes) is necessary, whereas investigations into transmission of genotypes and subtypes and identifying sources of infection and risk factors requires more discriminatory fingerprinting techniques, which can identify individual isolates or clonal lineages (Cacciò et al., 2005; Smith et al., 2006). Typing and subtyping systems used for human and nonhuman samples should also be used for environmental samples, particularly for source and disease tracking, in order to avoid any confusion arising from using different systems for human, nonhuman hosts, and environmental samples for veterinary and public health investigation of disease outbreaks (Cacciò et al., 2005; Smith et al., 2006). Species determination should be based on the analysis of at least two loci because this provides more robust information. At least one locus should be 18S rRNA and, where possible, another should be suitable for both species identification and further subtyping analysis. For Cryptosporidium, mini- and microsatellite typing and GP60 sequencing and analysis of a double-stranded RNA element have been used to subtype C. parvum and C. hominis, and might offer sufficient subspecies discrimination to address veterinary and public health investigations, either separately or in combination (reviewed in Cacciò et al., 2005; Smith et al., 2006). Recent work has confirmed the utility of GP60 sequencing and mini- and microsatellite markers in the study of the population structure of Cryptosporidium, and in understanding transmission dynamics of infection (Enemark et al., 2002; Xiao et al., 2004; Cacciò et al., 2005; Smith et al., 2006). The most commonly used PCR assays for detecting and typing Cryptosporidium are listed in Table 6.10. TABLE 6.10 List of Targets, Type of Assay and Main Use of Amplification-Based Techniques for Cryptosporidium Amplification Target 18S rDNA Hsp70 COWP Actin β-tubulin GP60 Microsatellites Minisatellites Extrachromosomal double-stranded RNA
Assay Type
Main Application
PCR, nested PCR, sequencing, PCR-RFLP, real-time PCR, microarray PCR, nested PCR, sequencing, real-time PCR, microarray PCR, nested PCR, sequencing, PCR-RFLP, microarray PCR, nested PCR, sequencing, PCR, nested PCR, sequencing, PCR-RFLP PCR, nested PCR, sequencing, PCR, nested PCR, sequencing, fragment typing PCR, nested PCR, sequencing, fragment typing Reverse transcriptase, PCR, sequencing, heteroduplex mobility assays
Species and genotype identification Species and genotype identification Species and genotype identification Species and genotype identification Species and genotype identification Subgenotype identification Subgenotype identification Subgenotype identification Subgenotype identification
Note: RFLP = restriction fragment length polymorphism; Hsp70 = heat shock protein 70; COWP = Cryptosporidium oocyst wall protein; GP60 = glycoprotein 60.
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Reporting Results of PCR-RFLP/Sequencing Examination
Negative results should be reported as “NO Cryptosporidium DNA detected.” Positive results should be reported as “Cryptosporidium DNA detected,” inserting the species/genotypes identified after identifying the respective species/genotypes from the RFLP profiles presented in Tables 6.7–6.9 or Cryptosporidium sequences deposited in GeneBank.
XIII. Shipping of Oocysts and Oocyst DNA for Quality Assurance and Round Robin Testing Exchange of oocysts and oocyst DNA among diagnostic laboratories for quality assurance and round robin testing should be encouraged to validate protocols for oocyst identification, DNA extraction, primer selection, and amplification and identification using PCR-based methods. Human infectious oocysts are Biosafety Level II organisms, and shipping must conform to IATA Dangerous Goods regulations (http://www.iata.org/ps/publications/9065.htm). Oocysts can be shipped air-dried onto microscope slides (either stained or unstained) or dried onto filter paper (e.g., Whatman FDA cards; http://www.whatman. com/products/?pageID=7.31.31), and oocyst DNA can be extracted in the recipient laboratory. One benefit of using slides containing stained oocysts is that they can be enumerated in both donor and recipient laboratories, thus adding an extra level of quality assurance to the system. Oocyst DNA can also be applied onto filter paper, dried, and then shipped. An import permit is generally required, but regulations differ from country to country, and laboratories should investigate these prior to shipment.
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McLauchlin, J., Amar, C.F.L., Pedraza-Diaz S., Mieli-Vergani, G., Hadzic, N., and Davies, E.G. 2003. Polymerase chain reaction-based diagnosis of infection with Cryptosporidium in children with primary immunodeficiencies. Pediatr. Infect. Dis. J. 22, 329–334. McLauchlin, J., Casemore, D.P., Harrison, T.G., Gerson, P.J., Samuel, D., and Taylor, A.G. 1987. Identification of Cryptosporidium oocysts by monoclonal antibody. Lancet (i), 51. Mead, J.R., Arrowood, M.J., and Sterling, C.R. 1988. Antigens of Cryptosporidium sporozoites recognized by immune sera of infected animals and humans. J. Parasitol. 74, 135–143. Meisel, J.L., Perera, D.R., Meligro, C., and Rubin, C.E. 1976. Overwhelming watery diarrhoea associated with Cryptosporidium in an immunosuppressed patient. Gastroenterology 70, 1156–1160. Morse, T.D., Nichols, R.A.B., Grimason, A.M., Campbell, B.M., Tembo, K.C., and Smith, H.V. 2007. Incidence of cryptosporidiosis species in paediatric patients in Malawi. Epidemiol. Infect. (in press). Moss, D.M., Bennett, S.N., Arrowood, M.J., Wahlquist, S.P., and Lammie, P.J. 1998. Enzyme-linked immunoelectrotransfer blot analysis of a cryptosporidiosis outbreak on a United States Coast Guard cutter. Am. J. Trop. Med. Hyg. 58, 110–118. Nichols, R.A.B. and Smith, H.V. 2004. Optimisation of DNA extraction and molecular detection of Cryptosporidium parvum oocysts in natural mineral water sources. J. Food Protect. 67, 524–532. Nichols, R.A.B., Campbell B.M., and Smith H.V. 2003. Identification of Cryptosporidium spp. oocysts in U.K. still natural mineral waters and drinking waters using a modified nested PCR-RFLP assay. Appl. Environ. Microbiol. 69, 4183–4189. Nichols, R.A.B., Moore, J.E., and Smith, H.V. 2006a. A rapid method for extracting oocyst DNA from Cryptosporidium positive human feces for outbreak investigations. J. Microbiol. Methods 65, 512–524. Nichols, R.A.B., Campbell B.M., and Smith H.V. 2006b. Molecular fingerprinting of Cryptosporidium species oocysts isolated during water monitoring. Appl. Environ. Microbiol. 72, 5428–5435. Nime, F.A., Burek, J.D., Page, D.L., Holscher, M.A., and Yardley, J.H. 1976. Acute enterocolitis in a human being infected with the protozoan Cryptosporidium. Gastroenterology 70, 592–598. Ong, C.S., Li, A.S., Priest, J.W., Copes, R., Khan, M., Fyfe, M.W., Marion, S.A., Roberts, J.M., Lammie, P.J., and Isaac-Renton, J.L. 2005. Enzyme immunoassay of Cryptosporidium-specific immunoglobulin G antibodies to assess longitudinal infection trends in six communities in British Columbia, Canada. Am. J. Trop. Med. Hyg., 73, 288–295. Ongerth, J.E. and Stibbs, H.H. 1987. Identification of Cryptosporidium oocysts in river water. Appl. Environ. Microbiol. 53, 672–676. Palmer, C.J., Xiao, L., Terashima, A., Guerra, H., Gotuzzo, H., Saldías, G., Bonilla, J.A., Zhou, L., Lindquist, A., and Upton, S.J. 2003. Cryptosporidium muris, a rodent pathogen, recovered from a human in Peru. Emerg. Infect. Dis. 9, 1174–1176. Perryman, L.E., Jasmer, D.P., Riggs, M.W., Bohnet. S.G., McGuire, T.C., and Arrowood, M.J. 1996. A cloned gene of Cryptosporidium parvum encodes neutralization-sensitive epitopes. Mol. Biochem. Parasitol. 80, 13–147. Pitlik, S.D., Fainstein, V., Garza, D., Guarda, L., Bolivar, R., Rios, A., Hopfer, R.L., and Mansell, P.A. 1983. Human cryptosporidiosis: spectrum of disease. Report of six cases and review of the literature. Arch. Intern. Med. 143, 2269–2275. Pohlenz, J., Moon, H.W., Cheville, N.F., and Bemrick, W.J. 1978. Cryptosporidiosis as a probable factor in neonatal diarhea of calves. J. Am. Vet. Assoc. 172, 452–457. Priest, J.W., Kwon, J.P., Moss, D.M., Roberts, J.M., Arrowood, M.J., Dworkin, M.S., Juranek, D.D., and Lammie, P.J. 1999. Detection by enzyme immunoassay of serum immunoglobulin G antibodies that recognize specific Cryptosporidium parvum antigens. J. Clin. Microbiol. 37, 1385–1392. Priest, J.W., Li, A., Khan, M., Arrowood, M.J., Lammie, P.J., Ong, C.S., Roberts, J.M., and Isaac-Renton, J.L. 2001. Enzyme immunoassay detection of antigen-specific immunoglobulin G antibodies in longitudinal serum samples from patients with cryptosporidiosis. Clin. Diagn. Lab. Immunol. 8, 415–423. Priest, J.W., Bern, C., Xiao, L., Roberts, J.M., Kwon, J.P., Lescano, A.G., Checkley, W., Cabrera, L., Moss, D.M., Arrowood, M.J., Sterling, C.R., Gilman, R.H., and Lammie, P.J. 2006. Longitudinal analysis of Cryptosporidium species-specific immunoglobulin G antibody responses in Peruvian children. Clin. Vaccine Immunol. 13, 139–144. Reed, C., Sturbaum, G.D., Hoover, P.J., and Sterling, C.R. 2002. Cryptosporidium parvum mixed genotypes detected by PCR-restriction fragment length polymorphism analysis. Appl. Environ. Microbiol. 68, 427–429.
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7 Immune Responses
Vincent McDonald
CONTENTS I. II.
III.
IV.
V.
Introduction .................................................................................................................................. 210 Innate Immunity ........................................................................................................................... 210 A. Components of Innate Immunity.................................................................................... 210 B. Epithelial Cells ................................................................................................................ 211 1. Toll-Like Receptors and NF-κB ...................................................................... 211 2. Production of Inflammatory Molecules........................................................... 211 3. Antimicrobial Peptides..................................................................................... 211 4. Complement ..................................................................................................... 212 C. Natural Killer Cells ......................................................................................................... 212 1. The Role of IFN-γ............................................................................................ 212 2. NK Cell Cytotoxicity....................................................................................... 213 T-Cell-Mediated Immunity........................................................................................................... 213 A. The Importance of T-Cells in the Control of Infection.................................................. 213 1. T-Cell Receptor αβ+ and γδ+ Cells .................................................................. 214 2. CD4+ and CD8+ T-Cells................................................................................... 214 B. T-Cells in Different Mucosal Compartments ................................................................. 215 1. Peyer’s Patches................................................................................................. 215 2. Lamina Propria................................................................................................. 216 3. Intraepithelial Lymphocytes............................................................................. 216 C. Parasite Antigen-Specific Priming of T-Cells and T-Cell Memory ............................... 216 Cytokines and Control of Infection ............................................................................................. 217 A. Th1 Cytokines ................................................................................................................. 217 1. IFN-γ ................................................................................................................ 217 2. IL-12................................................................................................................. 218 3. IL-2................................................................................................................... 219 B. Th2 Cytokines ................................................................................................................. 219 1. IL-4................................................................................................................... 219 2. IL-13................................................................................................................. 220 3. IL-5................................................................................................................... 220 C. Other Proinflammatory Cytokines .................................................................................. 220 1. TNF-α .............................................................................................................. 220 2. IL-1 and IL-6 ................................................................................................... 221 3. IL-18 and IL-15 ............................................................................................... 221 D. Cytokine-Mediated Parasite Killing ............................................................................... 221 Parasite-Specific Antibodies......................................................................................................... 222 A. Protective Role for Infection-Induced Antibodies.......................................................... 222 B. Protection with Antibodies Developed by Immunization .............................................. 223 1. Polyclonal Antibodies Obtained by Immunization with Crude Antigens ...... 223 2. Polyclonal Antibodies Obtained by Immunization with Recombinant Antigen or Parasite DNA................................................................................. 224 3. Antibodies from Immunized Chickens............................................................ 224 4. Passive Immunization with Monoclonal Antibodies....................................... 224
209
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VI. Vaccination Against Infection ...................................................................................................... 225 VII. Concluding Remarks .................................................................................................................... 225 References........................................................................................................................... 226
I.
Introduction
In 1984, as cryptosporidiosis was emerging as an important diarrheal disease of humans and domestic livestock, two landmark studies of Cryptosporidium were published. The first was the earliest report of mechanisms of protective immunity against the parasite using mice infected C. parvum (Heine et al., 1984) and the second was the description of an in vitro technique for growing this parasite in epithelial cells (Current and Haynes, 1984). These in vivo and in vitro infection models have since been the principal tools employed in the development of the Cryptosporidium immunology field, although neither model is ideal. The most important parasite species, C. parvum, can usually be cultured in epithelial cells only for a few days. Significantly, mice and other animals generally develop a poorly defined “natural resistance” to C. parvum infection before weaning (Sherwood et al., 1982), and C. hominis isolates may not normally develop in commonly used laboratory animal species (Morgan-Ryan et al., 2002). To circumvent natural resistance to C. parvum, neonatal or immunocompromised mice are often used for infection. Infections may also readily occur in transgenic mice with mutations affecting critical immune functions, and infection models involving these mice have been developed. Cryptosporidium muris has also been utilized and has the advantage of being able to infect adult animals, but it is not pathogenic and its development is restricted to the gastric epithelium (Iseki et al., 1989). The scope of immunological studies involving humans and livestock (mainly cattle and pigs) has been obviously more limited but, in general, most of the findings are in agreement with those obtained in murine studies. Considerable advances have been made in the understanding of the host responses to Cryptosporidium, including immune effector mechanisms that control infection, although there are still important gaps in our knowledge. This review will consider the nature of innate and adaptive immunity against Cryptosporidium in different host types. Examination of innate immunity will include the roles of epithelial cells, Toll-like receptors, NK cells, antimicrobial peptides, chemokines, and cytokines. Coverage of the adaptive immune responses will include assessments of the significance in immunity of CD4+ and CD8+ T-cells and cell-mediated Th1 versus humoral Th2 responses. Also, the involvement of B-cells and antibodies in control of infection will be evaluated, and the prospects for active and passive immunization will be considered.
II. A.
Innate Immunity Components of Innate Immunity
Innate immunity serves as an early sensor of infection and also activates evolutionarily conserved antimicrobial killing mechanisms that might curb the reproduction of invading microorganisms until the adaptive immune response becomes functional. The innate immune system includes the activities of inflammatory immune cells such as natural killer cells that produce cytokines and might be cytotoxic to infected, stressed, or tumor cells; macrophages and neutrophils that can engulf extracellular microorganisms; and eosinophils. Products released by these cells including mediators of inflammation, complement components and antimicrobial peptides are also important. Nonimmune cells, including epithelial cells, also have immunological functions such as the ability to produce cytokines and antimicrobial peptides. An important function of the adaptive immune system when activated by specific antigen recognition is to amplify elements of the innate immune response including antimicrobial killing mechanisms.
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Epithelial Cells
Epithelial cells are the only cells known to be infected by Cryptosporidium in vivo. Microarray analyses of 12,600 genes in a human enterocyte cell line infected with C. parvum showed that 223 were regulated by the parasite, including some involved in immune responses (Deng et al., 2004). Epithelial cells are now known to play a key part in establishing mucosal immune responses to infections. They act as sentinels and respond to infection by producing inflammatory molecules including chemokines involved in establishing inflammation. Enterocytes also generate mechanisms of microbial inactivation that can be activated directly by infection or via inflammation.
1.
Toll-Like Receptors and NF-κB
Epithelial cells express Toll-like receptors (TLRs), a group of innate molecular sensors of infection that ligate with conserved antigenic structures such as bacterial lipopolysaccharide (TLR4), bacterial cell wall components (TLR2), or DNA (TLR9) (Abreu et al., 2005). Cell signaling via an adaptor molecular that associates with TLRs, MyD88, leads to activation and migration to the cell nucleus of NF-κB, a key transcription factor for a variety of inflammatory molecules. NF-κB has been shown to be activated during C. parvum infection, and a key function for this molecule appears to be to prevent the infected cell undergoing apoptosis (Chen et al., 2001). It was shown that activation of a bile duct epithelial cell line via TLR2 and TLR4 occurs during C. parvum infection. Transfection of the cells with dominantnegative TLR2, TLR4, or MyD88 mutants or treatment of cells with interference RNA to the TLRs inhibited NF-κB activation by the parasite (Chen et al., 2005). MyD88 deficiency in these epithelial cells also increased parasite replication and in another study MyD881 mice had enhanced susceptibility to C. parvum infection (note that the MyD88 deficiency of these animals is not confined to epithelial cells) (Rogers et al., 2006). Treatment of neonatal mice with an oligodeoxynucleotide that is a ligand for TLR9 induced resistance to parasite reproduction (Barrier et al., 2006). Clearly, therefore, TLRs play a role in innate immunity, but the parasite molecules that act as ligands have yet to be identified.
2.
Production of Inflammatory Molecules
NF-κB promotes expression of proinflammatory molecules, including prostaglandins and chemokines. Indeed, C. parvum infection of epithelial cells induced PGE2, which stimulates production of mucin and regulates T-cell responses (Laurent et al., 1998). Chemokines produced by enterocytes play an important part in recruitment of immune cells into the intestine. Infection of cultures of mouse or human cell lines with C. parvum activated expression of numerous chemokines, including IL-8, GRO-α, RANTES, MCP-1, and MIP-2α, although variation between cell lines in the spectrum of chemokines was obtained (Laurent et al., 1997; Lacroix-Lamande et al., 2002). Inhibition of NF-κB activity in cells infected with C. parvum led to a decrease in IL-8 secretion (Chen et al., 2001). Immune cells recruited into the gut during infection include lymphocytes, macrophages, and neutrophils, and the functions of these cells during infection are of interest. Neutrophils may not be important as one study demonstrated that antibody-mediated depletion of neutrophils in C. parvum-infected piglets had no effect on either parasite reproduction or pathophysiology (Zadrozny et al., 2006).
3.
Antimicrobial Peptides
Intestinal epithelial cells are capable of producing antimicrobial peptides that can kill bacteria or parasites by disrupting their cell membranes. In addition, some peptides such as β-defensins have been shown to play a role in the development of adaptive immune responses (Bowdish et al., 2006). Raised expression of β-defensin was demonstrated in the intestine of C. parvum-infected calves (Tarver et al., 1998). It was shown that infection of mice or cell lines with the parasite regulated expression of β-defensins (Zaalouk et al., 2004). For example, human β-defensin 2 expression was upregulated by infection, whereas mouse β-defensin 1 was down-regulated. Importantly, human β-defensin 1 and -2 peptides killed sporozoites or prevented them from developing in vitro. It was subsequently found that TLR ligation was required for β-defensin expression by the parasite, and inhibition of TLR activity resulted in increased parasite reproduction in cell lines (Chen et al., 2005).
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212 4.
Cryptosporidium and Cryptosporidiosis, Second Edition Complement
Some complement components may be produced by enterocytes (Moon et al., 1997) but the role of complement in control of cryptosporidial infection is unclear. Classical activation of the complement cascade can be triggered by two mechanisms that lead to enzymic cleavage of C4 and C2 to form C3 convertase. Cleavage may be initiated by enzymatic action of C1 induced by antibody–antigen complexes or by serine proteases associated with mannose-binding lectin (MBL) following attachment of the latter to microorganisms. HIV patients with homozygous mutations in the MBL gene have been reported to be more susceptible to cryptosporidial infection (Kelly et al., 2000). MBL deficiency in the serum of Haitian children was also found to correlate with increased incidence of cryptosporidiosis (Kirkpatrick et al, 2006). Both MBL and C4 have been shown to adhere to the surface of sporozoites (Kelly et al., 2000), so it is possible that MBL may block parasite attachment to epithelial cells or activate the complement membrane attack complex.
C.
Natural Killer Cells
Natural killer (NK) cells are non-T, non-B lymphocytes that play a major role in the innate immune response against intracellular infection by viruses, bacteria, and parasites, and also tumor cells (Lodoen and Lanier, 2006). The two main mechanisms by which NK cells elicit antimicrobial activity are cytotoxic attack on infected cells and production of proinflammatory cytokines that stimulate antimicrobial killing mechanisms by other cells at the site of infection. Dendritic cells and macrophages are stimulated by infection to produce cytokines that activate NK cells. IL-12 is particularly important in this respect and may act synergistically with other proinflammatory cytokines such as IL-18 and TNF-α. A key cytokine produced by NK cells for control of intracellular infection is IFN-γ.
1.
The Role of IFN-γ
Studies of C. parvum infections in immunocompromised nude mice that are T-cell deficient and with SCID mice that are T- and B-cell deficient but have normal NK cell function indicated that IFN-γmediated innate immunity plays an important part in the control of the parasite’s reproduction. Adult immunocompromised mice develop chronic infection that is initially mild but becomes progressive and often fatal (Ungar et al., 1990a; Mead et al., 1991a; McDonald et al., 1992). Administration of anti-IFNγ-neutralizing antibodies to SCID or nude mice exacerbated infection and decreased the time taken for morbidity to occur, indicating the significance of IFN-γ in T-cell-independent immunity (Ungar et al., 1991; Chen et al., 1993b; McDonald and Bancroft, 1994). Similarly, IFN-γ-deficient SCID mice developed more intense infections than control SCID mice (Hayward et al., 2000). IL-12 plays an important part in this response to the parasite because treatment of newborn SCID mice with IL-12 prior to infection conferred resistance to C. parvum infection, whereas administration of anti-IL-12 neutralizing antibodies increased the level of infection (Urban et al., 1996). This is clearly a potent mechanism that helps to limit parasite reproduction for several weeks, but why the infection eventually overwhelms the host is unclear. Identification of NK cells as the source of IFN-γ in the mice, however, has been problematic. Different laboratories failed to observe any effect on the course of infection in immunocompromised mice after administration of antiasialoGM1 antibodies, which is a standard approach for depleting NK cells in mice (Ungar et al., 1991; McDonald and Bancroft, 1994). Similarly, treatment of SCID mice with an amount of anti-TNF-α antibodies that exacerbated systemic Listeria monocytogenes infection in the same host had no effect on C. parvum infection (McDonald and Bancroft, 1994). Also, administration of IL-2, which stimulates NK cell maturation, had no effect on the parasite’s reproductive capacity. In vitro studies, however, showed clearly that NK cells were likely to be the main source of IFN-γ. SCID mouse splenocytes produced IFN-γ in the presence of parasite antigen, and this effect was abrogated by depletion of NK cells and was inhibited by anti-IL-12- or anti-TNF-α-neutralizing antibodies (McDonald et al., 2000).
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NK Cell Cytotoxicity
NK cells display on their surface a complex array of activation and inhibitory receptors that regulate cytotoxicity (Lodoen and Lanier, 2006). NK cell surface inhibitory receptors recognizing the presence of MHC class I molecules on nucleated cells prevents development of lytic activity by the NK cells. If MHC class I expression is decreased, as happens during certain virus infections, the NK cells become cytotoxic and release the proteins perforin and granzyme, which mediate cell killing. Activating receptors induce cytotoxicity on contact with specific ligand molecules expressed by infected or stressed cells such as MHC-like MICA and MICB in humans, or pathogen-associated peptides expressed on the surface of infected cells (Lodoen and Lanier, 2006). Two studies suggest that cytotoxic activity by NK cells may also have a protective role. In the first, it was demonstrated that mice carrying the beige mutation that causes a defect in NK cell cytotoxic function (Roder and Duwe, 1979) had heavier C. parvum infections than mice without the mutation (Enriquez and Sterling, 1991). In the second, it was shown that NK cells from human blood had increased cytotoxicity against a human enterocyte cell line when infected with C. parvum (Dann et al., 2005). This activity was increased when the NK cells were incubated with IL-15, which is involved in the maturation and activation of NK cells. It was shown that the infected cell line had increased expression of the stress-associated molecule MICA, which induces NK cell cytotoxicity. In addition, ileal epithelium from a patient infected with the parasite had marked expression of MICA, whereas tissue from an uninfected patient did not (Dann et al., 2005).
III. T-Cell-Mediated Immunity The adaptive immune response induced by specific antigens recognized by T-cells and B-cells is generally required to eliminate rapidly proliferating or virulent microbial pathogens, and has the added advantage over innate immunity in having immunological memory, which allows prompt reactivation of memory T- and B-cells if reinfection occurs (Seder and Ahmed, 2003; Kalia et al., 2006). CD4+ T-cells become activated when in contact with antigen presenting cells such as dendritic cells and macrophages, and normally orchestrate the nature of adaptive immunity. T-cells activate B-cells to proliferate and differentiate into plasma cells that produce antibodies (Kalia et al., 2006).
A.
The Importance of T-Cells in the Control of Infection
Infection by Cryptosporidium leads to an intestinal inflammatory response involving lymphocytes and phagocytic cells (Tzipori, 1988). There is often villous atrophy and crypt hyperplasia, which is characteristic of T-cell-induced pathology (MacDonald and Spencer, 1992). Infection with C. parvum leads to a significant increase in different T-cell subsets in mucosal tissue (Boher et al., 1994; Abrahamsen et al., 1997). Also, there is increased expression of proinflammatory cytokines by intestinal T-cells that are associated with intestinal pathology (White et al., 2000; Lacroix et al., 2001). As infection is brought under control, the size of the T-cell population in the gut decreases to a normal level (Abrahamsen et al., 1997). There is, therefore, an associated T-cell activation, intestinal pathology, and control of infection. The first direct evidence that T-cells were essential for elimination of the parasite came from a study of C. parvum infection in athymic nude mice. Whereas wild-type neonatal animals had an acute infection that was readily cleared, T-cell deficient nude mice had chronic infection that was sometimes fatal (Heine et al., 1984). Chronic patterns of C. parvum and C. muris infection have been observed in adult nude and SCID mice (Ungar et al., 1990a; Mead et al., 1991a; McDonald et al., 1992), and depletion of T-cells in immunocompetent mice prevented recovery from C. muris infection (McDonald et al., 1992). Injection of nude or SCID mice with histocompatible lymphocytes brought about elimination of C. parvum infection, but if T-cells were removed from the donor cells, there was no protective effect (Mead et al., 1991b; Chen et al., 1993a; Perryman et al. 1994; McDonald and Bancroft, 1994).
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Relatively little work has been done to investigate the protective role of T-cells in other host species. In rats there was increased susceptibility to C. parvum infection if the animals were athymic (Gardner et al., 1991) or had been treated with immunosuppressive drugs (Rasmussen et al., 1991). In cattle, development of immunity is associated with activation of intestinal T-cells (Wyatt et al., 1997; Pasquali et al., 1997). Humans who underwent immunosuppressive chemotherapies or became immunosuppressed from viral infection had a higher than normal incidence of cryptosporidiosis; infections persisted unless immunocompetent status was restored (Tzipori, 1988). In humans, HIV infection can predispose to infection with Cryptosporidium, and individuals with active cryptosporidiosis and HIV infection T-cells in peripheral blood mononuclear cells (PBMC) were less able to proliferate and produce IFN-γ in the presence of cryptosporidial antigen than those from immunocompetent hosts (Gomez Morales et al., 1999). X-linked hyper-IgM syndrome is caused by a mutation in CD40L, a costimulatory molecule of T-cells that induces B-cell differentiation and IL-12 production by antigen presenting cells, and it has been shown that there are defects in T-cell effector functions in individuals with this mutation (Jain et al., 1999). The syndrome is linked with a high frequency of chronic cryptosporidiosis, and mice deficient either in CD40L or its ligand CD40 also develop severe disease after infection (Cosyns et al., 1998). Activation of T-cells may contribute to the pathogenesis of cryptosporidial infection, but T-cell deficiency leads to worsening of infections, indicating that these cells are involved in the establishment of immune effector mechanisms that eliminate the parasite.
T-Cell Receptor αβ+ and γδ+ Cells
1.
T-cells possess one of two types of heterodimer T-cell receptor (TCR), αβ or γδ. The TCR αβ has enormous structural diversity within the T-cell population, although each T-cell expresses only one form. A T-cell expressing TCR αβ may become activated when the receptor recognizes specific peptide antigen structures presented by MHC class II molecules on professional antigen-presenting cells, such as dendritic cells, or MHC class I molecules expressed on nucleated cells (Gao et al., 2002). γδ+ T cells form only 5–10% of T-cells but are relatively common within epithelium (Hayday and Tigelaar, 2003; Born et al., 2006). The γδ receptor has highly restricted structural variation and may recognize peptide or nonpeptide antigen ligands or even MHC-like stress molecules. Normally, αβ+ T-cells have a major involvement in immunity to infection compared with γδ+ T-cells. The functions of γδ+ T-cells are still relatively obscure, but they might be more diverse, having been associated with adaptive immunity, innate immunity, antigen presentation to other T-cells, and also downregulation of inflammation (Born et al., 2006). The role of TCR αβ+ and γδ+ cells in resistance to C. parvum infection was investigated using transgenic mice lacking one of the TCR types. Mutant mice lacking αβ+ T-cells (TCRα1) infected at 1 week of age were still infected 7 weeks later, whereas wild-type mice had cleared the infection after 3 weeks (Waters and Harp, 1996). When infected as adults, 60–70% of αβ+ T-cell-deficient mice were still shedding oocysts 12 weeks later, whereas most wild-type mice had no patent infection. Adult γδ+ T-celldeficient (TCRδ1) mice were as resistant to infection as wild-type mice, but when mice were infected as neonates, oocyst shedding lasted longer in the mutant animals. In another study, it was shown that C. parvum-infected TCRα1 mice had increased numbers of γδ+ T-cells in the intestinal intraepithelial lymphocyte population and that treatment with antibodies to deplete the γδ+ T-cells increased the level of infection (Eichelberger et al., 2000). In calves infected with C. parvum, both αβ+ and γδ+ T-cell numbers increased in the intestine as the infection developed and then declined as recovery progressed (Abrahamsen et al., 1997). These findings, therefore, imply that αβ+ T-cells are required for development of sterile immunity, whereas γδ+ T-cells, although not essential, do have a protective role that may be more significant in neonatal animals.
CD4+ and CD8+ T-Cells
2.
The αβ T-cells have different functions, depending on whether they express on their surface either CD4 (cytokine producing T helper cells) or CD8 (cytotoxic T-cells). CD4+ T-cells are activated by peptide antigen presented by MHC class II molecules on antigen-presenting cells, whereas CD8+ T-cells recognize peptides presented by MHC class I molecules (Gao et al., 2002). CD4 and CD8 bind to their +
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respective MHC molecules to optimize TCR-MHC interaction for T-cell activation. Investigations of CD4+ T-cell involvement in immunity to Cryptosporidium indicate that these cells are essential for control of infection. This was evident in a study with MHC class II-deficient mice that lack CD4+ Tcells, as they were unable to clear C. parvum infection for at least 8 weeks, whereas infections in wildtype mice had been cleared by this time (Aguirre et al., 1994). Similarly, C. parvum or C. muris infections were exacerbated in immunocompetent mice treated with anti-CD4 antibodies to deplete CD4+ T-cells (Ungar et al, 1991; McDonald et al., 1994). Injection of SCID mice with lymphocytes from immunocompetent animals allows them to resist cryptosporidial infection, but when CD4+ T-cells were depleted from the donor cells, the protective effect was lost (Chen et al., 1993a; McDonald et al., 1994). In humans with late-stage HIV infection and low CD4+ T-cell counts, there was increased susceptibility to cryptosporidial infection and the severity of disease was greater (Blanshard et al., 1992; Flanigan et al., 1992). After antiretroviral therapy and recovery of the CD4+ T-cell levels, cryptosporidial infection was readily cleared (Schmidt et al., 2001). It is evident, therefore, that CD4+ T-cells are major effector cells in immunity to cryptosporidial infection. The significance of CD8+ T-cells in immunity is perhaps less clear. Some investigations suggest that these cells are not required for effective immunological control of infection. In experiments with MHC class I-deficient mice that lack CD8+ T-cells, the mutant mice had similar low levels of C. parvum infection to wild-type mice after 4 weeks (Aguirre et al., 1994). In another study, reconstitution of SCID mice with spleen cells from immunocompetent mice depleted of CD8+ cells did not diminish the ability of mice to eliminate this parasite (Chen et al., 1993a). However, in a similar study depletion of the CD8+ T-cells from donor spleen cells delayed the recovery of the SCID mice (McDonald and Bancroft, 1994). Treatment of immunocompetent mice with anti-CD8 plus anti-CD4 antibodies increased C. parvum reproduction in mice more than anti-CD4 antibodies alone, although anti-CD8 antibodies alone had no effect on infection (Ungar et al., 1990a). In a C. muris investigation, administration of anti-CD8 antibodies enhanced parasite reproduction in mice, but they were not as effective as anti-CD4 antibodies (McDonald et al., 1994). Interestingly, during the first 24 h of C. parvum infection in mice, a degree of control associated with cytokine production could be demonstrated and was associated with αβ+ CD8+ T-cells (Leav et al., 2005). Overall, therefore, it is likely that CD8+ T-cells are involved in the development of adaptive immunity but have a subsidiary role to CD4+ T-cells and are not crucial.
B.
T-Cells in Different Mucosal Compartments
T-cell activities in three intestinal sites may be important in mucosal immune responses to infection (Mowat, 2003): (1) the organized lymphoid follicles such as Peyer’s patches in the small intestine, (2) the lamina propria underlying the epithelium, and (3) the epithelium itself where intraepithelial lymphocytes are found. Studies have been made to examine the importance of each of these mucosal compartments in response to cryptosporidial infection.
1.
Peyer’s Patches
Adaptive immune responses in the intestine may be initiated in the Peyer’s patches (PPs). Within the epithelium overlying mucosal lymphoid follicles are specialized epithelial cells called M cells that transport particulate antigens from the gut lumen into the lymphoid tissue where they undergo phagocytosis by dendritic cells and can be presented to T-cells. Ultrastructural studies have shown Cryptosporidium entering guinea-pig PP via M cells and the parasites were also observed within phagocytic cells underlying the epithelium (Marcial and Madara, 1986). Also, during C. parvum infection of mice, a 10-fold increase in numbers of certain T-cell subsets and B cells within PP has been reported (Mariotte et al., 2004). These findings imply that the PPs are key sites for initiating adaptive immune responses. Parasite antigen has also been detected in the mesenteric lymph nodes (MLN) that drain the intestine (Ponnuraj and Hayward, 2001) and are believed to be another site for initiation of mucosal T-cell responses. There is evidence that dendritic cells originating in the intestinal lamina propria can transport epithelial cells to the MLN (Huang et al., 2000). It is possible, therefore, that this is an alternative trafficking mechanism for bringing parasite antigen into contact with naïve T-cells.
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216 2.
Cryptosporidium and Cryptosporidiosis, Second Edition Lamina Propria
In the lamina propria (LP) there is significant recruitment of lymphocytes during C. parvum infection (Tzipori, 1988). In calves, increases in all the major subpopulations of T-cells were reported within 72 h of infection (Abrahamsen et al., 1997). Lymphocytes activated in PP migrate via the MLN and circulation, and are able to re-emerge in the intestinal LP as they express the integrin α4β7. Endothelial cells in the LP express the ligand for α4β7, the address in MAdCAM1. Early during infection 71 neonatal mice were shown to have increased numbers of parasites compared with wild-type mice, indicating that lymphocyte homing to the gut is important for establishing a protective response (Mancassola et al., 2004). In C. parvum-infected mice, the inflammatory response correlated with increased expression of chemokines by epithelial cells that attract T-cells from the local circulation into the LP (Lacroix-Lamande et al., 2002). Recovery from intractable cryptosporidial diarrhea associated with AIDS following highly active antiretroviral therapy was shown to coincide with a rapid increase in numbers of CD4+ T-cells in the LP (Schmidt et al., 2001).
3.
Intraepithelial Lymphocytes
Some lymphocytes in the LP-expressing αEβ7 move into the epithelium to form the intraepithelial lymphocytes (IELs), adhering to the basolateral surface of enterocytes via the ligand for αEβ7, E-cadherin (Mowat, 2003). IELs contain mainly T-cells, and those expressing CD8 or TCRγδ are more numerous than those with the CD4 marker (McDonald, 1999). The functions of these cells are not entirely clear as they are difficult to activate in vitro, but in an infection setting, IELs can produce proinflammatory cytokines and CD8+ IEL can demonstrate MHC class I-restricted cytotoxicity against infected epithelial cells. However, IELs can also have a regulatory role in ameliorating inflammation during infection, and γδ+ T cells might be important in this activity (Hayday and Tigelaar, 2003). As cryptosporidial infection is confined to the epithelium, it is important to understand the role of IELs in the host response, and evidence suggests the T-cells within this population can play a major part in the development of immunity. An increase in number of γδ+ cells in the IEL population of C. parvuminfected mice has been reported (Eichelberger et al., 2000). In calves, larger numbers of both CD4+ and CD8+ T-cells in the IEL population were found at the height of infection (Fayer et al., 1998) and around the time of recovery (Wyatt et al., 1999). In reinfected cattle there was an influx of CD8+ T-cells into the epithelium (Abrahamsen et al., 1997). It has been shown that SCID mice injected with IELs from immunocompetent mice previously infected by C. muris were able to control infection by this parasite (McDonald et al., 1996). Examination of the T-cell subpopulations present in the IELs isolated from the recovered SCID mouse cell recipients indicated the presence of CD4+ T-cells but there were larger numbers of CD8+ T-cells. However, depletion of either CD4+ or CD8+ cells from the donor population demonstrated that the transfer of immunity was associated only with CD4+ T-cells (McDonald et al., 1996). In a similar study with C. parvum, it was shown that immunity to infection could be adoptively transferred to SCID mice with infection-primed IELs, but the protective subpopulation was not identified (Adjei et al., 2000).
C.
Parasite Antigen-Specific Priming of T-Cells and T-Cell Memory
Many studies have shown that infection with C. parvum or C. muris induces parasite antigen-specific responses in T-cells. Spleen cells, MLN cells, or PBMC obtained from mice or humans infected with Cryptosporidium and cultured with oocyst antigen demonstrated an induction of T-cell proliferation and cytokine production (Harp et al., 1994; Tilley et al., 1995; Gomez Morales et al., 1999). During C. muris infection, the highest level of proliferation occurred around the time of parasite clearance (Tilley at al., 1995). Proliferation of spleen cells from mice previously infected with C. parvum involved mainly CD4+ T-cells, whereas little proliferation of CD8+ T-cells was obtained (Harp et al., 1994). This is in agreement with the view that CD4+ T-cells are more important for clearance of infection. Some studies, particularly involving C. muris, have measured the protective effect of lymphocytes from infection-primed animals after transfer to histocompatible-susceptible hosts. It was shown that cells from the spleen, MLN, or IEL from mice recovered from infection were significantly more effective in protecting SCID mice
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against the parasite than cells from uninfected donors (McDonald et al., 1994; McDonald et al., 1996). A similar protective effect was obtained against C. parvum using IEL from immune mice (Adjei et al., 2000). In another anomalous report involving C. parvum, however, it was shown that reconstitution of SCID mice with T-cells from ovalbumin-specific TCR transgenic mice conferred resistance against infection, suggesting that the protective T-cells did not have to be parasite antigen-specific (Lukin et al., 2000). Because adult mice do not develop the refractoriness to C. muris infection obtained with C. parvum, it has been possible to study immunological memory against this parasite. A primary infection induced strong resistance to challenge with oocysts given more than 30 d after recovery, whereas age-matched control mice were highly susceptible to challenge (McDonald et al., 1992). In other studies immunity was adoptively transferred to SCID mice using spleen cells from donors infected 97 d previously (McDonald et al., 1994). The protective effect of the donor cells lay mainly with the CD4+ T-cells, but infection-primed CD8+ T-cells were also beneficial. Immunity was also transferred using IEL isolated from donor mice 8 months after infection but these cells were less protective than ones obtained 3 months after infection, indicating that T-cell memory in response to Cryptosporidium wanes with time (McDonald et al., 2000).
IV.
Cytokines and Control of Infection
Depending on the nature of the antigens that the immune system encounters, CD4+ T helper (Th) cells may induce a cell-mediated immune response (Th1) or antibody-mediated response (Th2). These diverse Th responses are determined by the spectrum of cytokines produced by the T-cells themselves and by the antigen-presenting cells. In a Th1 response, IL-12 produced by dendritic cells and macrophages drives the T-cells to produce IFN-γ. This type of response is usually required to control and eliminate intracellular infections. In a Th1 response, CD8+ cytotoxic T-cells may also be induced with or without help from CD4+ T cells. These cells kill infected cells in an MHC class I-restricted fashion. A Th2 response is associated with production of IL-4, IL-5, IL-9, and IL-13, and polarized Th2 responses are observed in allergies and asthma; they may also be required to eliminate helminth infections (Wynn, 2003). The Th1 and Th2 pathways can oppose each other as Th1 responses are inhibited by Th2 cytokines such as IL-4 and IL-13 and, similarly, Th2 development is inhibited by Th1 cytokines. Recently, another Th response associated with inflammation has been characterized. Th17 cells are induced by the IL-12related cytokine IL-23, and they produce IL-17, IL-6, and TNF-α (Weaver et al., 2006). This response occurs in autoimmune diseases, but its involvement in immunity to infection is less clear.
A. 1.
Th1 Cytokines IFN-γγ
Most studies have indicated that the most effective adaptive immune response to Cryptosporidium infection involves IFN-γ activity. Treatment of immunocompetent mice, or SCID mice reconstituted with splenic T cells, with anti-IFN-γ-neutralizing antibodies increased reproduction of C. parvum and C. muris (Ungar et al., 1991, McDonald et al., 1992). However, repeated antibody administration did not prevent recovery from infection, suggesting IFN-γ-independent protective responses may also be acquired. In more definitive experiments it was shown that C57BL/6 IFN-γ1 mice developed rapidly growing fulminant C. parvum infections, but BALB/c IFN-γ1 mice were able to survive the infection (Theodos, 1998; Mead and You, 1998). The inability of C57BL/6 IFN-γ1 mice to control infection was associated with a defect in recruitment of T-cells and macrophages into the gut (Lacroix-Lamande et al., 2002). Infection with C. parvum has been shown to induce IFN-γ mRNA and protein expression in the intestine measured by RT-PCR and ELISA, respectively (Urban et al., 1996; Kapel et al., 1996). It has been observed in neonatal mice that the kinetics of IFN-γ expression reflects the pattern of acute infection, with the levels of IFN-γ increasing as infection approaches its peak level and declining rapidly as recovery
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gets under way (Kapel et al. 1996; McDonald et al., 2004). Stimulation of spleen cells from previously infected mice with oocyst antigen-induced proliferation of CD4+ T-cells and production of IFN-γ by these cells (Harp et al., 1994; Tilley et al., 1995). Further evidence of an association between CD4+ Tcells, IFN-γ, and immunity was suggested by the observation that treatment of mice with anti-CD4- plus anti-IFN-γ antibodies produced a greater exacerbation of C. parvum infection than either antibody alone (Ungar et al., 1991). Also, the ability of SCID mice to control C. muris infection after adoptive transfer of IEL required both the presence of CD4+ T-cells and IFN-γ activity (McDonald et al., 1996; Culshaw et al., 1997). However, a study of C. parvum in mice suggested that IFN-γ produced by CD8+ T-cells in the IEL population reduced parasite reproduction during the initial stage of infection (Leav et al., 2005). In calves, recovery from infection has been associated with increased expression of IFN-γ by IEL and LP cells (Fayer et al., 1998; Wyatt et al., 2001). In human studies, recovery from cryptosporidiosis in 15 patients correlated with PBMC producing IFN-γ when stimulated with parasite antigen (Gomez Morales et al., 1999). Severe chronic cryptosporidiosis in an HIV-negative child was closely associated with a deficiency in IFN-γ production (Gomez Morales et al., 1996). In another investigation with infected volunteers, however, IFN-γ expression detected in jejunal biopsies by in situ hybridization was observed predominantly in individuals with serological evidence of previous exposure to the parasite (White et al., 2000), but biopsies were obtained only once or twice and often late during infection. Some studies have measured the therapeutic effect of IFN-γ treatment on C. parvum development. In two studies involving mice, no protective effect was observed (McDonald and Bancroft, 1994; Kuhls et al., 1994), but administration of the cytokine to immunosuppressed rats was reported to reduce parasite numbers (Rehg, 1996). Also, remission of chronic cryptosporidiosis in a child was observed after treatment with IFN-γ (Gooi, 1994). It is likely that IFN-γ activity is necessary for early control of infection, therefore, but IFN-γindependent mechanisms of immunity also exist and in mice might act mainly later during a primary infection.
2.
IL-12
The early involvement of IL-12 in protection against C. parvum was suggested by two observations in the same study: firstly, treatment of immunocompetent neonatal mice with IL-12 prior to oocyst challenge, but not 24 h later, stimulated resistance to infection; secondly, administration of anti-IL-12neutralizing antibodies increased susceptibility to infection (Urban et al., 1996). Furthermore, an association between IL-12 activity and IFN-γ expression was established by the finding that IL-12 treatment increased the levels of IFN-γ mRNA in the intestine. In another study, strong expression of IL-12 mRNA in the intestines of neonatal BALB/c mice during C. parvum infection was allied to early control of infection (McDonald et al., 2004). That the protective role for IL-12 is closely linked with IFN-γ production was substantiated by the finding that treatment of IFN-γ1 mice with IL-12 did not enhance immunity to C. parvum and, paradoxically, daily administration of IL-12 exacerbated the infection and reduced the level of parasite antigen-specific responses of spleen cells (Smith et al., 2001). IL-12 is composed of two subunits p40 and p35, whereas the related cytokine IL-23 which is important in Th17 inflammatory responses, comprises p40 and p19. Adult IL-12p401 mice that are deficient in both IL-12 and IL-23 on a C57BL/6 background were highly susceptible to C. parvum infection but recovered and were resistant to reinfection (Ehigiator et al., 2003). In the absence of IL-12, these mice were still able to mount an IFN-γ response (Ehigiator et al. 2005), suggesting that other cytokines can replace IL-12 for the induction of IFN-γ production although they are not as effective as IL-12. Whereas C57BL/6 mice and neonatal BALB/c mice require IL-12 for efficient control of infection, adult BALB/c showed little dependence on IL-12 because p401 mice, similar to wild-type mice, developed very mild infections (Campbell et al., 2002). Further investigation with C57BL/6 transgenic mice by the same group demonstrated that IL-12p351 mice, deficient only in IL-12, were only slightly less susceptible than IL-12p401 mice to infection, implying that IL-12 was more important than IL-23 in development of immunity to C. parvum (Ehigiator et al., 2005). Hence, Th17-mediated inflammation might have a relatively minor involvement in protective immunity in mice.
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In C. parvum-infected calves, the IEL and LP cell populations showed increased levels of IL-12 mRNA (Fayer et al., 1998). The effect of treatment of cattle with recombinant IL-12 on infection was different from that observed in neonatal mice (see above). Administration of the cytokine on one occasion prior to infection or on several days around the time of oocyst challenge did not affect parasite reproduction, although there was an increase in the number of intestinal T-cells and also in expression of IFN-γ (Pasquali et al., 2006). IL-12, therefore, is expressed while a Th1 response develops during a primary C. parvum infection and in mice plays an important part in inducing IFN-γ expression required for early parasite clearance. Other cytokines may also be involved in driving the Th1 response, however, and in mice the degree of dependence on IL-12 for development of resistance may differ between strains.
3.
IL-2
IL-2 has been considered as a Th1 cytokine and plays an important part in the induction of NK cell and T-cell proliferation. PBMC from C. parvum- infected patients and spleen cells from C. muris-infected mice produced large quantities of IL-2 as well as IFN-γ when stimulated with oocysts antigen (Gomez Morales et al., 1999; Tilley et al., 1995). Treatment of mice with anti-IL-2 neutralizing antibodies, however, had no effect on susceptibility to infection (Ungar et al., 1991; Enriquez and Sterling, 1993) and treatment of mice with exogenous IL-2 did not influence parasite reproduction (McDonald and Bancroft, 1994). These findings suggest that IL-2 is not an essential cytokine for development of immunity.
B. 1.
Th2 Cytokines IL-4
Investigations of the involvement of IL-4 in immunity have produced conflicting conclusions. Reports suggest IL-4 has either no role in immunity, inhibits development of immunity, promotes the protective Th1 response early in infection, or is involved late during infection in parasite clearance in an IFN-γindependent manner. In a study comparing C. muris reproduction in MHC congenic strains of mouse, it was found that BALB/b mice (H-2b) had prolonged infections compared with BALB/c mice (H-2d) and the increased susceptibility of the H-2b mice correlated with a high level of production of IL-4 by spleen cells early during infection (Davami et al., 1997). BALB/c mice, in contrast, produced IL-4 only late during infection when parasite numbers were in steep decline (Davami et al., 1997; Tilley et al., 1995). Another investigation showed that adult C57BL/6 mice had few intestinal IL-4-producing CD4+ Tcells early during infection, but large numbers of these cells appeared during the late part of infection when parasites were being eliminated (Aguirre et al., 1998). A protective role for the IL-4-producing cells was implied by the observation that IL-41 mice of this strain or mice treated with anti-IL-4 antibodies had extended infections compared with controls (Aguirre et al., 1998). Also, the IL-4 involvement in immunity appeared to be IFN-γ-independent as anti-IFN-γ-neutralizing antibodies administered late in infection had no effect on the recovery process. However, another group could find no increased susceptibility of adult C57BL/6 IL-41 mice to C. parvum infection (Campbell et al., 2002). In yet another study, when BALB/c neonatal mice were treated with anti-IL-4 antibodies they produced greater numbers of C. parvum oocysts around the time of the peak of infection than control mice (McDonald et al., 2004). Similarly, neonatal BALB/c IL-41 mice were more susceptible to infection than wild-type mice but, significantly, both groups recovered at the same time. IL-4 protein was detected by Western blotting in the intestines of wild-type mice 24 h postinfection; treatment of wild-type mice with IL-4 prior to infection, but not on day 4, increased resistance to infection (McDonald et al., 2004). These findings suggested that in neonatal BALB/c mice IL-4 was required to establish the early control of infection. The source of IL-4 was not determined and its mechanism of action is unclear. However, IL-41 mice had lower levels of intestinal IL-12 and IFN-γ than wild-type mice at the peak of infection, indicating IL-4 could promote the Th1 response. IL-4 has been shown to increase IL-12 production by dendritic cells (Hochrein et al., 2000) and stimulate maturation of IFN-γ-producing T cells (Noble and Kemeny,
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1995). IL-4 also acted synergistically with IFN-γ to activate antimicrobial killing by intestinal epithelial cells (Lean et al., 2003). In models of Th1-mediated diseases involving BALB/c mice, treatment with IL-4 exacerbated colitis (Fort et al., 2001) and uveitis (Ramanathan et al., 1996) providing further evidence that IL-4 can enhance IFN-γ-mediated responses. In studies of IL-4 expression during C. parvum infection of humans, it was demonstrated by in situ hybridization and immunocytochemistry that IL-4 is expressed in the gut of C. parvum-infected persons if they had previously been exposed to the infection (Robinson et al., 2001a). There was no correlation between symptoms or oocyst excretion levels and expression of the cytokine. In infected cattle, measurement of IL-4 mRNA in intestinal cells showed little expression around the time of recovery (Wyatt et al., 2001).
2.
IL-13
IL-13 has overlapping functions with IL-4, and the two cytokines employ the same receptor, IL-4Rα (Wynn, 2003). BALB/c IL-4Rα1 mice, like the IL-41 mice, were found to have more intense C. parvum infection than wild-type mice and the rate of recovery was initially slower than in IL-41 mice (McDonald et al., 2004). This suggests, therefore, that IL-13 may have a protective role in this mouse strain. IL-13, unlike IL-4, did not increase the capacity of IFN-γ to inhibit C. parvum reproduction in an enterocyte cell line, however (Lean et al., 2003). Malnourished Haitian children with persistent cryptosporidiosis were found to have a marked intestinal inflammatory response and expressed IL-13 but not IFN-γ (Kirkpatrick et al., 2002), which does not signify a protective role for the cytokine.
3.
IL-5
The first study of the role of Th2 cytokines in control of cryptosporidial infection compared the requirement for IL-4 and IL-5 in immunity by administration of cytokine-neutralizing antibodies to mice infected with C. parvum (Enriquez and Sterling, 1993). The most effective antibody for increasing parasite replication was anti-IL-5, whereas a combination of anti-IL-5 and anti-IL-4 antibodies was more effective than anti-IL-5 alone. The results suggested that IL-5 was important, and IL-4 had a protective role in concert with IL-5. It has been reported that IL-5 is expressed during infection of BALB/c IFNγ1 mice that recover from infection (Bonafonte et al., 2000), but no further investigation has been made of the protective role of this cytokine in resistance to infection. Clearly, further studies of IL-4, and other Th2 cytokines, including IL-5 and IL-13, are required to elucidate their contribution to the protective host response against Cryptosporidium in different hosts.
C. 1.
Other Proinflammatory Cytokines α TNF-α
TNF-α is a proinflammatory cytokine that is commonly produced by T-cells and other cells during infections and is important in the stimulation of antimicrobial killing mechanisms as well as in inflammation. Numerous studies have shown that this cytokine is up-regulated in the intestine during C. parvum infection of mice (Lacroix et al., 2001; Ehigiator et al., 2005), humans (Robinson et al., 2001b) and in infected human intestinal xenografts in SCID mice (Seydel et al., 1998). The failure of C57BL/6 IFNγ1 mice to control infection correlated with poor intestinal TNF-α expression compared with infected wild-type mice, suggesting that the cytokine may have an important protective effect (Lacroix et al., 2001). This view was supported by the additional finding that treatment of these mice with TNF-α reduced parasite reproduction (Lacroix et al., 2001). In addition, TNF-α was shown to significantly inhibit C. parvum development in human and mouse enterocyte cell lines (Pollok et al., 2001; Lean et al., 2006). However, other investigations indicated that TNF-α activity was not necessary for the host control of infection. In separate studies it was reported that administration of anti-TNF-α neutralizing antibodies failed to increase C. parvum reproduction in mice (McDonald et al., 1992; Chen et al., 1993b). Furthermore, no
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differences in levels of infection were obtained between TNF-α1 and wild-type mice (Lean et al., 2006). Hence, at best TNF-α may have a redundant role in establishing resistance to infection.
2.
IL-1 and IL-6
It is possible that other proinflammatory cytokines such as IL-1 and IL-6 that are produced by a variety of cell types may provide similar activities in the absence of TNF-α. IL-1 may be up-regulated during infection of mice and humans (Lean et al., 2006; Robinson et al., 2001b), and this cytokine has been shown to inhibit parasite reproduction in enterocytes (Pollok et al., 2001). IL-6 expression was increased in C. parvum-infected mice with Th1 deficiencies (Lacroix et al., 2001; Ehigiator et al., 2005) and reduced parasite development in vitro (V. McDonald, unpublished data).
3.
IL-18 and IL-15
IL-18 and IL-15 are produced by macrophages, dendritic cells, and also epithelial cells, and activate both NK cells and T cells to produce IFN-γ. IL-15 is also important for the homeostasis of NK cells and memory T cells. There was increased IL-18 mRNA expression in the intestines of C. parvum-infected IL-12/IL-23-deficient mice (Ehigiator et al., 2005) and enhanced IL-18 mRNA and protein expression in human enterocyte cell lines infected by the parasite (McDonald et al., 2006). Also, exogenous IL-18 reduced multiplication of the parasite in these cell lines, suggesting a possible autocrine protective effect for this cytokine (McDonald et al., 2006). Immunocompetent volunteers infected with C. parvum who had symptoms but failed to express IFNγ in the intestine usually expressed IL-15 (Robinson et al., 2001a). In cattle, however, infection resulted in reduced expression of IL-15 (Canals et al., 1998). In IL-12-deficient C57BL/6 mice that develop heavy infections but are able to establish production of IFN-γ and recover, IL-15 is expressed early in infection (Ehigiator et al., 2005) and so might be important in inducing Th1-mediated immunity in the absence of IL-12.
D.
Cytokine-Mediated Parasite Killing
An important function of IFN-γ and other proinflammatory cytokines is to activate antimicrobial killing mechanisms that include production of toxic nitric oxide derivatives or oxygen radicals, or creating a deficiency of metabolites required for growth of microorganisms such as tryptophan or cellular iron (Rottenberg et al., 2002). High levels of nitric oxide production can be stimulated by IFN-γ, often acting in concert with other cytokines such as TNF-α (Nacy et al., 1991). The cytokines induce expression of inducible nitric oxide synthase (iNOS) which converts arginine to citrulline and nitric oxide (Oswald et al., 1994). Cryptosporidium parvum infections grew slower in nude mice fed a diet supplemented with arginine than in mice fed a normal diet (Leitch and He, 1994), whereas treatment with an iNOS inhibitor, N-nitro-L-arginine, exacerbated infection. In further studies it was observed that plasma levels of the nitric oxide derivatives nitrate and nitrite increased during infection as did iNOS expression detected by immunohistochemistry (Leitch and He, 1999). In addition, infections were moderately exacerbated in iNOS1 mice and treatment of mice with a nitric oxide donor reduced oocyst and infection scores. Increased iNOS activity was also observed in the ileal mucosa of infected piglets, particularly in enterocytes, and treatment with an iNOS inhibitor increased the infection level but did not affect features of pathology (Gookin et al., 2006). An additional benefit of nitric oxide production was that it increased expression of PGE2 which enhanced the barrier function of the infected epithelial cells measured by electrical resistance in Ussing chambers (Gookin et al., 2004). Collectively, these findings support a role in parasite killing for nitric oxide in murine and porcine infections. In none of these studies was any attempt made to determine mechanisms by which iNOS expression/activity was upregulated. Two other murine studies found no involvement for nitric oxide in control of infection. Inhibition of nitric oxide production by treatment with an amount of aminoguanidine sufficient to exacerbate Listeria infection had no effect on C. parvum infection scores (Kuhls et al., 1994), while iNOS1 mice were found to have similar levels of infection compared with control animals (Hayward et al., 2000).
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Using an in vitro infection model, an investigation was made of possible mechanisms by which IFNγ inhibited C. parvum reproduction in human enterocyte cell lines (Pollok et al., 2001; Lean et al., 2003). The use of NOS inhibitors showed that nitric oxide production was not essential for parasite killing. Catabolism of tryptophan appeared not to be involved either as supplementing culture medium with an excess of the amino acid did not reduce IFN-γ activity. Two mechanisms of IFN-γ action were identified, however; the cytokine-treated enterocytes were moderately resistant to parasite invasion and the inhibition of intracellular parasite development was largely overcome by an excess of Fe2+ in the culture medium, indicating that deprivation of cellular Fe was an important killing mechanism (Pollok et al., 2001). Inhibition of development was also obtained with TNF-α and IL-1, but neither of these cytokines produced a synergistic killing effect with IFN-γ. TNF-α was shown to act mainly to prevent parasite invasion and appeared to have little effect on intracellular parasite development (Lean et al., 2006). IL4 had no effect on parasite development by itself but, surprisingly, acted synergistically with low concentrations of IFN-γ to prevent parasite replication (Lean et al., 2003). Therefore, in addition to promoting development of a Th1 response (McDonald et al., 2004), IL-4 can interact with IFN-γ at the effector stage of immunity for parasite elimination.
V.
Parasite-Specific Antibodies
Many studies have been made of the properties of antibodies generated as a result of cryptosporidial infection or immunization with antigen preparations. Investigations have attempted to ascertain the significance of infection-induced antibodies in clearing parasites. Other studies have examined whether colostral antibodies developed either through infection or immunization could provide passive immunity to offspring or other infected hosts. In addition to their role in protection, parasite-specific antibodies have been important for identifying immunodominant antigens and antigens involved in sporozoite attachment to host cells and invasion. Furthermore, monoclonal antibodies developed from immunized mice have been valuable for the development of diagnostic reagents and in technologies for detection of parasites in environmental samples.
A.
Protective Role for Infection-Induced Antibodies
Parasite-specific antibodies appear in the circulation and in the mucosa during infection of different mammalian hosts, including humans, cattle and sheep (Ungar et al., 1986; Peeters et al., 1992; Hill et al., 1990). IgM, IgG, and IgA titers measured by ELISA generally increase during infection and decline after recovery, although IgG in serum may persist for several months longer than IgM (Ungar et al., 1986). In a study with adult human volunteers, however, secretory IgA (sIgA) was detected in fecal samples during infection but neither IgG nor IgM was found (Dann et al., 2000). In developing countries parasite-specific serum IgG titers in children can continue to rise with time (Priest et al., 2006; Cox et al., 2005) presumably because of continual environmental exposure to the parasite but it was reported that in cases of persistent diarrhea IgA and IgM levels decreased (Khan et al., 2004). Infection antibodies have been shown to recognize a wide range of oocyst polypeptides from low to high molecular weights (Hill et al., 1990; Peeters et al., 1992). Secretory IgA (sIgA) produced by a host as a result of infection can play a major part in protecting the mucosal surface from toxins and microbial pathogens. In C. parvum-infected immunocompetent human adults the level of parasite-specific sIgA was higher in individuals excreting oocysts or with diarrhea (Dann et al., 2000). Patients with congenital hypogammaglobulinemia have been unable to clear C. parvum infections (Lasser et al., 1979), suggesting antibody is required for immunity. However, AIDS patients with chronic cryptosporidiosis were reported to have high titers of sIgA (Cozon et al., 1994), possibly signifying that additional mechanisms were necessary for control of infection. In contrast, another study showed that AIDS patients with a strong serological response to a 27-kDa parasite antigen had a reduced risk of cryptosporidial diarrhea (Frost et al., 2005). In rodent studies, passive protection against C. parvum infection was obtained in nude mice following oral administration of bile containing
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parasite-specific sIgA isolated from rats that had recovered from infection (Albert et al., 1994). Also, sIgA monoclonal antibodies recognizing the C. parvum surface antigen p23 prepared with PP B-cells from infected mice were able to provide a degree of protection to neonatal mice after passive transfer (Enriquez and Riggs, 1998). Investigations with B-cell-deficient mice, however, suggest antibodies play little part in elimination of Cryptosporidium. Following depletion of B-cells in neonatal mice by administration of anti-μ chain antibodies, it was observed that mice were no more susceptible to C. parvum infection than control mice (Taghi-Kilani et al., 1990). Similarly, C. muris infection in SCID mice reconstituted with spleen cells depleted of B-cells followed the same acute pattern as in mice receiving cells that were not depleted of B-cells (McDonald et al., 1992). Also, in a study with transgenic mice lacking B-cells (μMT1), the kinetics of infection and clearance were the same for mutant and control animals (Chen et al., 2003). There are also conflicting findings on the capacity of Cryptosporidium-specific antibodies passively acquired by offspring to protect against infection. Chicks born to hens that had been previously infected with C. baileyi were more resistant to the same infection than control chicks although the relationship between parasite-specific antibodies and resistance to infection was not measured (Hornok et al., 1998). Uninfected newborn calves were shown to have IgM, IgG, and IgA antibodies to the immunodominant C. parvum sporozoite surface antigen p23 in fecal samples and these may have come from colostrum (Wyatt et al., 2000). Oral administration to calves of oral bovine serum concentrate containing anti-C. parvum antibodies reduced the levels of diarrhea and parasite development after infection (Hunt et al., 2002). However, humans with chronic cryptosporidiosis failed to respond to oral administration of bovine colostrum containing anticryptosporidial antibodies (Saxon and Weinstein, 1987). In a murine study of lacteal immunity, neonatal pups of previously infected dams were not more resistant to C. parvum infection than control pups (Moon et al., 1988). Some human epidemiological studies suggest that breastfed babies are less likely to experience cryptosporidiosis (e.g., Molbak et al., 1994; Enriquez et al., 1997), but other investigations were not in agreement with this finding (Nchito et al., 1998). The often contradictory observations on the part played by infection antibodies in immunity probably suggest that they may have a limited capability to reduce parasite reproduction but are not a vital factor in host resistance.
B. 1.
Protection with Antibodies Developed by Immunization Polyclonal Antibodies Obtained by Immunization with Crude Antigens
The question of whether colostrum from a host previously exposed to cryptosporidial infection can passively transfer immunity to offspring may be unresolved, but more encouraging results have been obtained with colostrum from dams mucosally immunized with parasite material. In early studies, socalled hyperimmune bovine colostrum (HBC) was obtained after injection of crude C. parvum oocyst antigen preparations plus adjuvant into the mammary glands of preparturient cattle (Tzipori et al., 1986; Fayer et al., 1989a; Ungar et al., 1990b). Oral administration of HBC to humans with severe disease was shown, in some instances, to decrease the parasite burden and symptoms (Tzipori et al., 1986; Ungar et al., 1990b). Variable results might be attributed to the severity of disease or the immune status of the patients. However, in a study involving 16 healthy volunteers in which the prophylactic effect of HBC was examined, no significant beneficial effect was observed (Okhuysen et al., 1998). The results obtained with cattle were better because calves treated with HBC before infection had diarrhea for 0–4 days whereas for controls diarrhea lasted 4–6 days (Fayer et al., 1989a). The duration of oocyst excretion was usually shorter in the treated group of calves. Treatment of neonatal mice also resulted in a decreased level of infection (Fayer et al., 1989b). As HBC contained high titers of cryptosporidial antibodies, the possible role of antibodies in passive protection was also investigated. Incubation of sporozoites with HBC reduced their infectivity when transferred to neonatal mice (Fayer et al., 1989b). Infections in neonatal mice given HBC antibodies (IgA, IgM, IgG1, or IgG2) were less intense than in control mice (Fayer et al., 1990). Immunogold ultrastructural studies of infected mouse tissue showed that IgA, IgG, and IgM from HBC recognized
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meronts, merozoites, and gametes of C. parvum (Fayer et al., 1991), and in Western blots these antibodies reacted with a large number of oocyst and sporozoite antigens (Tilley et al., 1990).
2.
Polyclonal Antibodies Obtained by Immunization with Recombinant Antigen or Parasite DNA
Good protection from C. parvum infection was obtained in calves receiving colostrum from cows vaccinated subcutaneously with the recombinant p23 antigen (present in both sporozoites and merozoites) plus adjuvant (Perryman et al., 1999). When infected, all calves receiving control colostrum developed diarrhea whereas calves given colostrum from immunized dams had no diarrhea and a 99.8% reduction in oocyst production. Immunization with plasmid DNA encoding C. parvum antigens resulted in production of parasitespecific antibody. Following jet-injection of a plasmid encoding the sporozoite surface antigen CP15/60 into muscle or mammary glands of sheep, high titer antibodies to the CP15/60 antigen were obtained in serum and colostrum, although these titers were obtained more reliably with intramammary injection (Jenkins et al., 1995). Colostrum obtained after injection by the mammary route when transferred to drug-immunosuppressed mice reduced development of C. parvum by about 50% (Jenkins et al., 1999). Nasal immunization of pregnant goats with a plasmid containing CP15-DNA led to the suckling kids infected with C. parvum shedding fewer oocysts and having greater weight gains than infected kids from unvaccinated goats (Sagodira et al., 1999).
3.
Antibodies from Immunized Chickens
Chickens immunized with C. parvum antigens produced eggs that contained high titers of anti-C. parvum antibodies or antibodies that reduced binding of sporozoites to enterocytes, and when transferred to SCID mice, the antibodies provided partial protection against infection by the parasite (Cava and Sterling, 1991; Kobayashi et al., 2004). However, administration of C. baileyi antigens to the amniotic sac of chick embryos resulted in increased susceptibility of the hatched chick to infection with this parasite species (Hornok et al., 2000).
4.
Passive Immunization with Monoclonal Antibodies
Murine monoclonal antibodies (mAb) developed after immunization with C. parvum oocysts antigens have been shown by immunofluorescence microscopy to specifically recognize different structures of the oocysts including the wall, the whole sporozoite, the sporozoite surface, and the apical pole (McDonald et. al., 1991). When sporozoites were incubated with certain mAbs that recognized surface antigens, they became less infectious for neonatal mice in a time-dependent manner (Perryman et al., 1990). Three mAbs could cross-react with merozoites and incubation of antibody with isolated merozoites neutralized the capacity of the merozoites to infect mice (Bjorneby et al., 1990). Individual mAbs orally administered to neonatal mice prior to infection did not provide any passive immunity to infection, but a combination of three antibodies given on 3 consecutive days from the start of infection reduced the infection score (Arrowood et al., 1989). Another study investigated passive immunization of neonatal mice with three neutralizing mAbs that reacted with different sporozoite surface antigens—CSL, gp25200, and p23—either used individually or in combinations. Prophylactic treatment with one antibody or pairs of antibodies starting when oocysts were inoculated produced moderate resistance to infection, but a formulation containing all three antibodies produced additive protection amounting to a reduction in infection levels of 86–93% (Schaefer et al., 2000). A different result was obtained when these antibodies were used therapeutically in adult SCID mice with established chronic infections (Riggs et al., 2002). Only one of the mAbs (3E2) that reacts with the CSL antigen given alone was able to reduce parasite reproduction, and combinations with other antibodies did not improve efficacy. The level of extraintestinal infection in the bile ducts was also decreased by mAb 3E2, but infection in the pylorus was unaffected. It seems, therefore, that the efficacy of parasite-specific antibody may be dependent on the level of infection.
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VI. Vaccination Against Infection As chemotherapeutic compounds have limited success in the treatment of cryptosporidiosis, a plausible rationale for immunological investigation of Cryptosporidium has been to develop attenuated live or recombinant vaccines. Comparatively little research effort has been aimed directly at formulation of a vaccine to protect against infection, probably because there are considerable scientific obstacles, and there is also an issue of expediency. An effective method for attenuating C. parvum may be γ-irradiation of oocysts. Relatively low doses of irradiation (200 krad) did not affect excystation efficiency but no excystation occurred with a higher dose (5000 krad) (Kato et al., 2001). Sporozoites given 200 krad were able to invade epithelial cells in vitro but did not develop. Irradiation of oocysts also decreased the infectivity of oocysts for mice (Yu and Park, 2003). In another study a dose of irradiation was established that prevented parasite reproduction after oral administration of the oocysts to day-old calves but induced a degree of resistance against subsequent challenge with viable oocysts (Jenkins et al., 2004). As for subunit vaccination, many C. parvum antigens have been characterized (Okhuysen and Chappell, 2002) but none has been identified as suitable for vaccination. Problems that may arise with live vaccines are that they might be prohibitively expensive to manufacture, and a cold chain between production and use in the field may be an unacceptable logistical complication (Jenkins, 2001). In the case of subunit vaccines, little success has been achieved in developing highly effective protection against any protozoan infection (Jenkins, 2001). For Cryptosporidium immunization, it may be necessary to administer a subunit vaccine by the oral route to obtain a protective response in the gut, in which case oral tolerance that prevents inflammatory responses against harmless antigens has to be overcome (Mowat, 2003). New mucosal adjuvants may in the future help to overcome this problem, however (Moyle et al., 2004). Candidate vaccines need to be identified using an immunocompetent host that is highly susceptible to infection. In the case of the major parasite species of mammals, C. parvum, this is problematic as animals develop natural resistance to infection. Natural resistance also means that protection against infection is needed only during a relatively narrow window of time early in life, and the immature immune system of neonates may not respond well to vaccination. The approach of protecting young livestock by active immunization may not, therefore, be practicable. For humans, vaccines might only have relevance in areas of developing countries with poor hygiene, and it might be necessary to immunize against at least two species, C. parvum and C. hominis.
VII. Concluding Remarks Innate immunity may make an important contribution to the control of cryptosporidial infection. A prominent component in the innate immune response appears to be the infected epithelium that acts as a sentinel of infection, shapes the early local inflammatory response and directly inactivates parasites. NK cells are the likely source of IFN-γ produced independently of T-cells that provides a robust partial protection against infection in mice and may also have cytotoxic activity against infected cells. The T-cell response that is necessary to clear the infection is dependent on the CD4+ T-cell subpopulation with perhaps a moderate contribution by CD8+ T-cells. As with innate immunity T-cell-dependent control of parasite reproduction involves IFN-γ activity. IFN-γ and other proinflammatory cytokines can act directly on enterocytes to induce mechanisms for parasite inactivation. Studies suggest that there are also IFN-γ-independent immune mechanisms, but the nature of these is unknown. Findings on the role of Th2 cytokines in immunity are conflicting and, in the case of the most studied cytokine, IL-4, its involvement in immunity may depend on the age and genetic make-up of the host. Cryptosporidial infection leads to a parasite-specific antibody response in both serum and intestine but murine studies suggest that B cells are not essential for recovery from infection. Epidemiological studies suggest colostrum or parasite-specific colostral antibodies may provide a degree of passive immunity to suckling human offspring, but results from experimental studies with animals suggest
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previously infected dams do not protect offspring. There is evidence that a degree of passive immunity can be obtained with orally administered colostral antibodies from dams immunized mucosally with crude or recombinant oocyst antigens and adjuvant or DNA plasmids encoding parasite antigens. Whether this approach to immunotherapy can be efficiently scaled up and be cost effective remains to be seen.
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8 Clinical Disease and Pathology
Cirle Alcantara Warren and Richard L. Guerrant
CONTENTS I.
Clinical Manifestations ................................................................................................................ 235 A. The Immunocompetent Host........................................................................................... 235 B. The Immunocompromised Host ..................................................................................... 236 C. Age-Related Infections.................................................................................................... 237 Cryptosporidium Infection in Children ........................................................... 237 1. 2. Cryptosporidium Infection in the Elderly ....................................................... 238 II. Organ Sites ................................................................................................................................... 239 A. GI-Tract ........................................................................................................................... 239 B. Extraintestinal Sites......................................................................................................... 239 III. Histopathology ............................................................................................................................. 240 IV. Pathophysiology ........................................................................................................................... 241 V. Human Infectivity Trials .............................................................................................................. 242 A. Prepatent and Patent Periods .......................................................................................... 242 B. Infectious Dose................................................................................................................ 242 C. Clinical versus Subclinical Infections ............................................................................ 243 D. Postinfection Sequelae in Healthy Hosts........................................................................ 244 VI. Conclusion.................................................................................................................................... 244 References........................................................................................................................... 244
I.
Clinical Manifestations
Infection with Cryptosporidium has been documented in both immunocompetent and immunocompromised patient populations. Clinical manifestations vary according to the immune and nutritional status of the host. Clinical features of human cryptosporidiosis in different patient groups are summarized in Table 8.1.
A.
The Immunocompetent Host
The majority of cases in immunocompetent hosts are sporadic and can involve either children or adults, in both the developed and developing world (Navin, 1985). Infection has also been reported in animal handlers (Current et al., 1983; Fayer and Ungar, 1986), homosexual men (Soave et al., 1984), travelers (Jokipii et al., 1985, Soave and Ma, 1985), and contacts of infected individuals (for example, household contacts or hospital personnel) (Koch, 1985; Wolfson et al., 1985; Fayer and Ungar, 1986; Ribeiro and Plamer, 1986). The three major clinical presentations in immunocompetent persons are asymptomatic carriage, acute diarrhea (most common), and persistent diarrhea. In diarrheal illness, the onset is usually acute after an incubation of 3 to 14 days and is generally self-limiting. Infection results in gastrointestinal illness 235
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TABLE 8.1 Clinical Features of Cryptosporidiosis Immunocompetent Persons
Immunocompromised Persons
Age-Related Populations
Susceptible population
Healthy adults, human volunteers
Children, especially those less than 2 years of age, elderly adults
Site of infection Enteric presentation
Intestinal Asymptomatic, acute
Signs and symptoms Duration of illness
Diarrhea, abdominal discomfort, fatigue 4–14 days
Persons with AIDS, solid-organ transplants, malignancies, malnutrition, hemodialysis, primary immunodeficiency diseases Intestinal or extraintestinal Asymptomatic, transient, chronic, or fulminant Diarrhea, weight loss, abdominal pain, nausea, vomiting, fever 14 days to months
Outcome
Self-limiting
Characteristics
Associated with increased morbidity and mortality
Primarily intestinal Asymptomatic, acute, persistent Diarrhea, dehydration, malnutrition, weight loss 5–17 (outbreak) > 14 days to weeks in children Associated with long-term developmental impact on malnourished children, increased transmission and hospitalization in the elderly
characterized by malaise or fatigue, anorexia, vomiting, abdominal pain, and cramps in most patients. More than half the patients experience weight loss, nausea, flatulence, low-grade fevers (up to 38˚C), chills, sweats, myalgias, or headaches (MacKenzie et al., 1994; Mathieu et al., 2004; Bushen et al., 2006). The hallmark of infection is diarrhea, which is watery, voluminous, and occasionally explosive and foul smelling. Blood and pus are not present in the stool, and the fecal leukocyte examination is negative. Diarrhea usually lasts 6 to 14 days, although some patients have a more prolonged illness. Persons infected during the outbreak in Milwaukee had a mean duration of symptoms for 12 days and a maximal number of 12 stools per day (MacKenzie et al., 1994). Diarrhea recurred in more than a third of the visitors with laboratory-confirmed infection and in 21% of residents with clinical disease (MacKenzie et al., 1995). In some immunocompetent persons diarrhea can persist for a month, or rarely, as long as 4 months (Isaacs 1985; Wolfson et al., 1985; Jokipii and Jokipii, 1986; Soave and Armstrong, 1986; Stehr–Green et al., 1987).
B.
The Immunocompromised Host
Cryptosporidiosis is prevalent in human immunodeficiency virus (HIV)-positive patients, particularly those with a low CD4 count and those who engage in high-risk sexual practices (reviewed in Hunter and Nichols, 2002). Patients with malignancies, including those with hematological malignancies undergoing chemotherapy or bone marrow transplantation, patients with solid-organ transplants, and patients on hemodialysis, also have a high prevalence of cryptosporidiosis (Gentile et al., 1991; Tanyuksel et al., 1995; Sreedharan, et al., 1996; Turkcapar et al., 2002). Other immunocompromised hosts at risk of cryptosporidial infection are those with primary immunodeficiency diseases such as combined immunodeficiencies, antibody deficiencies, complement deficiencies, and defects in phagocyte number and function (reviewed by Hunter and Nichols, 2002). Diarrhea can occasionally be much more severe (for example, 17 L of daily stool output) in immunocompromised hosts with defects in either humoral or cell-mediated immunity (CDC, 1982; Navin and Juraneck, 1984; Fayer and Ungar, 1986). Diarrhea in patients with AIDS can be “fulminant,” with passage of more than 2 L of stool per day, or “cholera-like” (Blanshard et al., 1992; Manabe et al., 1998). Although asymptomatic carriage, transient infection, and self-limiting acute disease also occur, infection is frequently persistent or chronic and the resulting morbidity—as in AIDS patients—can contribute to earlier mortality (Navin and Hardy, 1987; Blanshard et al., 1992; Manabe et al., 1998). In the 1993 Milwaukee outbreak, the affected immunocompromised patients had more diarrheal stools per day (mean,
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15 versus 12; p = 0.08) and were more likely to be hospitalized than immunocompetent patients (odds ratio, 1.9; 95% CI, 0.95 to 3.9) (MacKenzie et al., 1994). Weight loss is common in chronic cryptosporidiosis. In an outbreak in Nevada in 1994, two-thirds of HIV-infected patients developed chronic disease and suffered a median weight loss of 13.6 kg (Goldstein et al., 1996). Over half of the affected patients died within 6 months of the onset of the outbreak, and 60% of those had cryptosporidiosis listed as a cause of death on their death certificate. In addition to developing a more severe intestinal disease, immunocompromised hosts can also develop atypical disease presentations in the gastrointestinal tract as well as extraintestinal sites, presumably via infection from the intestinal lumen, rather than by hematogenous or systemic dissemination. These unusual presentations are discussed in the following text.
C. 1.
Age-Related Infections Cryptosporidium Infection in Children
Cryptosporidium is a leading cause of persistent diarrhea in children in developing countries (Dillingham et al., 2002). Young children, particularly those less than 2 years of age, are more likely to be infected with Cryptosporidium than are older children (Perch et al., 2001; Kirkpatrick et al., 2002; Pereira et al., 2002; Kahn et al., 2004; Gatei et al., 2006). In a recent prospective case-control study in Bangladesh, the most common symptoms in children infected with Cryptosporidium were watery diarrhea and vomiting (Kahn et al., 2004). Similar to infection in adults, the total duration of diarrhea was greater in Cryptosporidium-infected patients than in controls who had diarrhea but without cryptosporidial infection (mean ± SD = 12 ± 4.8 days for cases versus 6 ± 2.9 days for controls), and 41% of the infected children presented with or subsequently developed persistent diarrhea compared with none of the uninfected controls.
a.
Day-Care Outbreaks
A review was conducted of 14 outbreaks of cryptosporidiosis in day-care centers in the United States, Europe, Australia, Chile, and South Africa from 1984 to 1992 (Cordell and Addiss, 1994). Although the overall prevalence in cross-sectional studies in day-care centers in the United States, Spain, and France ranged from 1.8 to 3.8%, the number of centers with one or more infected children ranged from 11.8 to 42.8%. The highest rates were found in non-toilet-trained toddlers and staff caring for diapered children, usually in late summer or early fall. In outbreaks in day-care centers, 4 to 23% of infected children might not have diarrhea, and in non-outbreak conditions 56 to 75% of oocyst-positive children were asymptomatic (Cordell and Addiss, 1994). Infectious oocysts can be excreted up to 5 weeks after diarrhea has stopped. Day-care attendance continues to be identified as a risk factor for cryptosporidiosis (Pereira et al., 2002; Craun and Calderon, 2006). In recent outbreaks in day-care centers in Brazil and Spain, attack rates ranged from 6.4 to 46% (Franco and Cordeiro, 1996; Rodriguez-Hernandez et al., 1996; Cordell et al., 1997; Carvalho-Almeida et al., 2006; Goncalves et al., 2006; Teresa Ortega et al., 2006). The highest rates were also found in children 2 years of age or less who wear diapers, who have concomitant illness, who have relatives with diarrhea, or when one relative is a worker at the day-care facility. In studies where genotyping was performed, C. hominis was the predominant organism isolated in the outbreaks (Goncalves et al., 2006; Teresa Ortega et al., 2006).
b.
Nutritional Impact
Malnourished children are at greater risk of infection and suffer greater consequences than well-nourished children. Stunting, wasting, or both have been observed in malnourished children with cryptosporidiosis (Macfarlane and Horner-Bryce, 1987; Sallon et al., 1988; Sarabia-Arce et al., 1990; Garcia Velarde et al., 1991; Sallon S et al., 1994; Bhattaacharya et al., 1997; Javier Enriquez et al., 1997; Banwat et al., 2003; Tumwine et al 2003; Hamedi et al., 2005). In a longitudinal study in Brazil, children born with low birth weight (less than 2500 g) had a greater than fivefold risk of having a first symptomatic episode of Cryptosporidium infection, compared with all other children (Newman et al., 1999). Although this finding suggests that prior poor nutritional status predisposes to children to subsequent infection, infection
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itself results in weight loss, which might not be regained later. Weight gain was impaired in the month following symptomatic and even asymptomatic cryptosporidial infections in children in Peru, where malnutrition did not appear to be a significant risk factor for infection (Checkley et al., 1997). Followup data from 185 children in the cohort revealed that those with cryptosporidial infection experienced several months of both weight and height faltering (Checkley et al., 1998). Children who were infected between birth and 5 months did not catch up in height and exhibited a deficit of almost 1 cm a year later, compared with uninfected children of the same age. Stunted children did not catch up in weight or height a year later compared to uninfected, stunted age-matched children. Similarly, in 1064 children in West Africa, cryptosporidiosis in infancy caused significant weight and height losses that persisted even at 2 years of age (Molbak et al., 1997). Cryptosporidiosis, especially during infancy, was associated with excessive and protracted (nearly 2 years) diarrheal disease burden, most likely contributing to malnourishment in young children (Agnew et al., 1998) and possibly impairment in physical fitness, cognitive development, and school performance several years later (Guerrant et al., 1999; Niehaus et al., 2002; Lorntz et al., 2006). Clinical illness, dehydration (even in this age of oral rehydration therapy), need for hospitalization, or subsequent mortality were increased in children with cryptosporidial infection, especially those with malnutrition or those with underlying immunocompromised status (Macfarlane and Horner-Bryce, 1987; Sarabia-Arce 1990; Molbak et al., 1993; Cicirello et al., 1997; Guarino et al., 1997; Amadi et al., 2001; Tumwine et al., 2003; Abdel Messih et al., 2005). Concomitant HIV infection increased the likelihood of malnourishment in Ugandan children with persistent diarrhea secondary to Cryptosporidium (Tumwine et al., 2005). Furthermore, shedding in favela children is greater even than in patients with AIDS (Bushen et al., 2007). Data on the impact of breast-feeding on the susceptibility to Cryptosporidium are less consistent than data on the interaction of nutritional status and cryptosporidial infection. Early reports suggested that bottle-feeding was a risk factor in acquiring cryptosporidiosis in children less than 18 months of age (Hojlyng et al., 1986) and that oocysts were rarely found in stools of infants receiving only breast milk (Pape et al., 1987). Most subsequent studies on the prevalence of cryptosporidiosis in children associated with breast-feeding, especially exclusive breast-feeding, found lower infection rates (Sallon et al., 1994; Javier Enriquez et al 1997; Bhattacharya et al., 1997; Kirkpatrick et al., 2002; Adjei et al., 2004; AbdelMessih et al., 2005). In a study specifically examining the impact of breast-feeding on children with diarrhea (not necessarily secondary to Cyptosporidium), the incidence of diarrhea was noted to be higher in weaned children compared with even partially breast-fed children in both 1- and 2-year olds (relative risks of 1.41, 95% CI of 1.23, 2.15; and 1.7, 95% CI of 1.29, 2.15; respectively) (Molbak et al., 1994). Duration of diarrhea was also longer by 1 day among weaned children and the mortality was 3.5 times higher among weaned children aged 12 to 35 months of age compared with breast-fed children. Whether similar effects can be claimed in cryptosporidial infection remains to be proven. The mechanism as to how breast-feeding can confer protection against Cryptosporidium is unclear. The levels of Cryptosporidium-specific IgA in breast milk of mothers do not correlate with the prevalence or duration of cyrptosporidial illness among Peruvian children (Sterling et al., 1991). Other epidemiologic studies do not show any association between breast-feeding and decreased susceptibility to the infection (Nchito et al., 1998; Pereira et al., 2002; Khan et al., 2004; Hamedi et al., 2005).
2.
Cryptosporidium Infection in the Elderly
Although the elderly are at higher risk for infections, including gastrointestinal infections, there are very little data on cryptosporidial infection among this population. Seroprevalence of cryptosporidiosis increases with age (Zu et al., 1994; Frost et al., 1998). Examination of sera from 1356 National Health and Nutrition Examination Survey (NHANES) III participants revealed strong responses to cryptosporidial 15/17-kDa antigen among older age groups, blacks, Hispanics, and low-income groups (Frost et al., 2004). The 15/17-kDa marker peaked at 4 to 6 weeks and remained elevated until 4 to 6 months postinfection (Priest et al., 2001; Muller et al., 2001). In the NHANES study, cryptosporidial IgG was detected in sera from 67.6% of persons 70 years of age and older. Although seropositivity might mean protection against clinical disease (Chappell et al., 1999), this finding also suggests susceptibility to infection, although mostly asymptomatic, among individuals of advanced age. In a small rural area in
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Korea where 77% of the population was 50 years of age or older, Cryptosporidium oocysts were found in 69% of 29 residents 60 to 69 years of age and in 54% of 20 residents 70 years of age or older (Chai et al., 2001). About half of the infected residents experienced only mild symptoms of diarrhea and intermittent abdominal discomfort, which lasted for 1 to 2 weeks; the rest were asymptomatic. Elderly patients with underlying diseases in hospitals or in nursing homes might be more susceptible to symptomatic infection and possibly increased mortality (Bannister and Mountford, 1989; Fripp et al., 1991; Lee et al., 2005; Pandak et al., 2006). Cryptosporidiosis in elderly Rhode Island residents was the cause of nosocomial diarrhea initially suspected of being caused by Clostridium difficile (Neill et al., 1996). In a hospital in India, Cryptosporidium was identified as the etiologic agent in 18.3% elderly patients with diarrhea; 17% presented with vomiting and abdominal pain and 31% presented with fever (Gambhir et al., 2003). The number of daily bowel movements ranged from 3 to 9, and the duration of diarrhea lasted 5 to 17 days. During the Milwaukee outbreak of 1993, the daily rates of gastroenteritis-related emergency room visits and hospitalizations among individuals 65 years of age or older were substantially higher than before the outbreak (Naumova et al., 2003). The median incubation period was also shorter in the elderly compared with younger adults (5 to 6 days versus 8 to 9 days). In addition, secondary spread appeared more pronounced in the elderly, suggesting a higher risk for secondary person-to-person transmission in this population, a finding that may be relevant among those residing in nursing homes, hospitals, or even in households (Neill et al., 1996; Naumova et al., 2003; Pandak et al., 2006). Cryptosporidiosis can also be an initial presentation for unsuspected HIV infection. Therefore, other coexisting immunocompromising conditions such as AIDS need to be considered in this population as well (Schoofs et al., 2004). The clinical features and consequences of cryptosporidial infection among the elderly need further investigations.
II.
Organ Sites
A.
GI-Tract
Cryptosporidium is primarily a pathogen of the small bowel causing blunting of the microvilli, submucosal edema, and mononuclear cell inflammatory infiltration in the lamina propia. Although the parasite tends to preferentially infect the jejunum and ileum, the colon can be heavily infected without infection of the small bowel. Multisite involvement has been described, and widespread infection of the intestinal tract, including the small and large bowels, or localized involvement of the proximal small intestines, has been associated with more severe diarrheal illness (Clayton et al., 1994; Lumadue et al., 1998). Clinically, intestinal involvement presents as diarrhea and is often associated with malabsorption, such as abnormal D–xylose tests and evidence of intestinal barrier disruption by lactulose–mannitol permeability ratios (Adams et al., 1994; Goodgame et al., 1995; Lima et al., 1997). Atypical gastrointestinal manifestations of cryptosporidiosis have also been reported. In the immunocompromised host, particularly those with AIDS, case reports of gastric involvement, which could be complicated by antral narrowing and pneumatosis cystoides intestinalis, possibly leading to bowel perforation, have been published (Garone et al., 1986; Cerosimo et al., 1992; Collins et al., 1992; Sidhu et al., 1994; Samson and Brown, 1996; Iribarren et al., 1997; Ventura et al., 1997; Moon et al., 1999; Clemente et al., 2000). Isolation of Cryptosporidium from gastric mucosa has been reported in about a third of patients with AIDS who underwent endoscopy for chronic diarrhea and/or unexplained gastrointestinal symptoms (Rossi et al., 1998). Most of these patients did not have symptoms attributable to the pathogen in the stomach. Esophageal involvement presenting as dysphagia and vomiting in a 2year-old child and appendicitis have been very rarely reported as well (Kazlow et al., 1986; Oberhuber et al., 1991; Buch et al., 2005).
B.
Extraintestinal Sites
The biliary tract has been a well-known site of cryptosporidial disease in patients with AIDS (Forbes et al., 1993; Lopez-Velez et al., 1995, Vakil et al., 1996; Chen and LaRusso, 2002), and rare cases have
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been reported in bone marrow and solid-organ transplant recipients and immunocompetent children (Campos et al., 2000; Goddard et al., 2000; Westrope and Acharya, 2001; Abdo et al., 2003; Dimicoli et al., 2003). Cryptosporidium is the most common cause of AIDS-cholangiopathy (Chen and LaRusso, 2002). Biliary involvement in AIDS patients is associated with low CD4 counts and increased mortality (Hashmey et al., 1997; Vakil et al., 1996). Radiographic imaging revealed dilation of ducts, thickening of gallbladder walls, and pericholecystic fluid collection (McCarty et al., 1989; Teixidor et al., 1991; Vakil et al., 1996). The clinical features usually include right upper quadrant abdominal pain, nausea, vomiting, and fever, accompanied by elevated serum alkaline phosphatase levels. Hepatobiliary disease can mimic pancreatic involvement (de Souza et al., 2004). However, involvement of the pancreas itself has been documented in autopsy cases (Godwin, 1991). Acute and chronic forms of pancreatitis have been described in cases presenting with severe abdominal pain, elevated serum amylase levels, abnormal radiologic findings, and endoscopic retrograde cholangiopancreatography revealing papillary stenosis (Calzetti et al., 1997). Cryptosporidium oocysts have been reported in respiratory secretions of patients with diarrhea and respiratory symptoms. Coexistence with other pathogens and asymptomatic presentation are common. In Spain, 16% (7/43) of patients with chronic diarrhea due to Cryptosporidium had oocysts detectable in the sputum (Lopez-Velez et al., 1995). Five of these patients had respiratory symptoms and abnormal chest radiographs. Mycobacterium tuberculosis was isolated from two of the five and M. avium was isolated from two others. The remaining two patients had no respiratory symptoms and normal chest radiographs. In another case series from Spain, five patients had apparent “pulmonary cryptosporidiosis.” Four of them had another pathogen isolated (M. tuberculosis in two, M. fortuitum in one, and cytomegalovirus and Pneumocystis in one) (Clavel et al., 1996). Although cases of biopsy-proven pulmonary cryptosporidiosis are rarely reported, the clinical significance of isolation of oocysts in the sputum remains unclear.
III. Histopathology In humans, detailed studies of intestinal histopathology and function are provided only in case reports of immunocompromised patients, primarily with AIDS. Infection is confined to the luminal border of the enterocytes (Orenstein, 1997), similar to what is demonstrated in vitro in human intestinal cell lines. The parasite burden and degree of injury vary from area to area. Ultrastructural studies generally show that the intestinal mucosa is intact and the enterocytes are well preserved (Lefkowitch et al., 1984; Modigliani et al., 1985). Microvilli are displaced at the sites of parasite attachment to the enterocyte surface and may be elongated next to the parasite. In addition, “peaking” or “pedestal formation” of the host cell may occur at the point of attachment of the cryptosporidia (see Chapter 1) (Vetterling et al., 1971; Lefkowitch et al ., 1984). In some infections, villous architecture by light microscopy is moderately to severely abnormal, revealing crypt elongation and villous atrophy, and may be accompanied by a mixed inflammatory-cell infiltrate in the lamina propria (Meisel et al., 1976 , Soave et al., 1984; Lumadue et al., 1998). In areas of severe injury, the tall columnar cells are replaced by distorted, disorganized cells undergoing degeneration and necrosis (Orenstein et al., 1997). Enterocytes often appear vesiculated with inconspicuous brush borders and may show blunted microvilli at higher magnification (see Chapter 1). Strips of sloughing cells can be seen. The degree and cell content of inflammatory infiltrate in the lamina propria in clinically affected patients has ranged from minimal (Soave and Armstrong, 1986) to substantial (Meisel et al., 1976; Modigliani et al., 1985; Soave et al., 1984) and may include plasma cells, lymphocytes, macrophages, and/or polymorphonuclear leukocytes. Interstitial edema accompanying the mixed inflammatory infiltrates has been observed (Godwin et al., 1991). Reactive epithelial changes and neutrophil infiltration have been described in gastric infection (Lumadue et al., 1998). Interestingly, paneth cells, which are known to secrete antimicrobial peptides and proteins, are decreased in intestinal tissues of patients diagnosed with cryptosporidiosis compared with other infective or inflammatory conditions (Kelly et al., 2004). Intracellular granules and defensin-5 secretion were also depleted during infection.
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Infection of the bile ducts and gallbladder also vary from area to area (Margulis, 1986; Kahn, 1987; Schneiderman, 1987; Teixidor et al., 1991; Chen and LaRusso, 2002). Edema and inflammation are more pronounced than in the intestinal tract. The hepatobiliary ducts are dilated with focal strictures, the walls of the hepatobiliary ducts and gallbladder are thickened, and the lumen is filled with necrotic debris, inflammatory cells, and parasites. Cryptosporidium has been isolated in both biopsy and cytology specimens in about half of the patients with biliary involvement (Vakil et al., 1996).
IV.
Pathophysiology
In contrast to some animal studies where the degree of pathologic abnormality has tended to correlate with the extent of infection and severity of clinical illness (Tzipori et al., 1980; Tzipori, 1983; Heine, 1984; Moon et al., 1985), in humans there can be significant diarrhea (≥ 1 L/day) with minimal histopathology in the gut (Modigliani et al., 1985). The mechanisms by which Cryptosporidium causes diarrhea in either the immunocompetent or the immunocompromised host have not been fully elucidated. Both secretory (unaffected by fasting) and malabsorptive diarrhea have been reported in AIDS patients infected with Cryptosporidium (Koch, 1983; Petras, 1983; Soave et al., 1984; Modigliani et al., 1985). D-xylose and vitamin B12 malabsorption, steatorrhea, and increased fecal alpha1-antitrypsin clearance have been reported (Adams et al., 1994; Clayton et al., 1994; Goodgame et al., 1995; Lima, 1997). Radiographic studies have also showed results consistent with malabsorption, such as flocculation of the barium, mucosal thickening, and dilation in the small bowel (Berk et al., 1984). Detailed electron micrograph studies indicate that Cryptosporidium develops intracellularly in the host epithelial cell but is extracytoplasmic (Vetterling et al., 1971; Current and Reese, 1986; Chapter 1). The attachment of the sporozoites, which is ligand–receptor mediated (reviewed in Smith et al., 2005; Chapter 3), triggers rearrangement of the host actin cytoskeleton to form a parasitophorous vacuole within which the parasite develops (Chen and LaRusso, 2000) and exerts its effects on the host and surrounding cells. Indeed, C. parvum infection of a human ileocecal cell line led to upregulation of multiple host genes for actin and tubulin and downregulation of genes for actin-binding proteins, confirming its role in modifying the host cytoskeletal structure (Deng et al., 2004; Chapter 3). The observation that voluminous, watery diarrhea can occur in the immunocompromised host might suggest an enterotoxin or neurohumoral product caused by the parasite. Experimental results are mixed (Casemore et al., 1985; Garza et al., 1986; Guerrant et al., 1990). Thorough investigations have failed to discover such a toxin or secretagogue (Sears and Guerrant, 1994; Guarino et al., 1994, 1995). Observations in humans with cryptosporidiosis and in experimental porcine infections suggest that part of the symptomatology is due to malabsorption secondary to blunted or inflamed villi coupled with intact or, possibly, enhanced fluid secretion from the crypts (Goodgame et al., 1995). In suckling rats, both amino acid absorption and Na-glucose transport were impaired during infection (Capet et al., 1999; Topouchian et al., 2001). Furthermore, malabsorption of amino acids seemed to persist beyond clearance of infection in the rats (Topouchian et al., 2003). The attachment of Cryptosporidium to epithelial cells activates NFΚB, which induces antiapoptotic mechanisms as well as upregulates proinflammatory cytokine/chemokine expressions (Chen et al., 2001). Cytokines, produced by either the enterocyte or recruited inflammatory cells in the lamina propia, may elicit prostaglandin-dependent secretion. TNFhas been shown to stimulate chloride secretion through a prostaglandin mediator (Guarino et al., 1994, 1995; Argenzio et al., 1993, 1994; Clark and Sears, 1996). Prostaglandin-independent net electrogenic transport across the ileal mucosa in C. parvum-infected suckling rat has also been described (Barbot et al., 2001). Interferon-γ has been implicated in the development of leaky and dysfunctional epithelium via its effects on the sodium/hydrogen ion exchangers and sodium–potassium-dependent ATPase (Rocha et al., 2001; Sugi et al., 2001). Indeed, the roles of cytokines and inflammatory cells are further confirmed by in vivo and human studies revealing detection of TNF-, IL-8, and lactoferrin (in malnourished children) during infection (Seydel et al., 1998; Robinson et al., 2001; Kirkpatrick et al., 2002, 2006; Alcantara et al., 2003; Chapter 7). Substance P (SP), a neuropeptide known to cause chloride secretion in the gastrointestinal tract, and SP-receptor mRNA were increased in jejunal samples from C. parvum-infected macaques. Elevated basal ion secretion and glucose malabsorption were also noted in the animals
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(Hernandez et al., 2006). These findings support previous findings, where SP mRNA and protein expression were higher in jejunal tissue samples from symptomatic patients with AIDS and cryptosporidiosis compared with immunocompetent volunteers with self-limited infection (Robinson et al., 2003). In vitro studies have demonstrated the destructive effects of Cryptosporidium, including loss of barrier integrity (increased transepithelial resistance) and epithelial cell injury. These observations are consistent with the increased lactulose-mannitol excretion in adult patients or children with cryptosporidiosis (Adams et al., 1994; Griffiths et al., 1994; Goodgame et al., 1995; Lima et al., 1997; Zhang et al., 2000). Direct cytopathic effects of C. parvum in biliary and intestinal cells (Chen et al., 1998; McCole et al., 2000) may be secondary to either caspase- and/or Fas/Fas ligand-dependent apoptosis (Ojcius et al., 1999; Chen et al., 1999). Activation of NFΚB in directly infected cells possibly limits apoptosis in uninfected adjacent intestinal and biliary cells, ensuring survival of the organism and persistence of infection (Chen et al., 2001). The molecular details as to how Cryptosporidium mediates epithelial cell apoptosis and secretion of proinflammatory cytokines are just beginning to be clarified. An analysis of the gene profile expression of the host epithelial cells infected with C. parvum indicates that genes for proinflammatory chemokines IL-8, RANTES, and SCYB5, in addition to genes for cytoskeletal structure, are upregulated during infection. Genes associated with cell proliferation and apoptosis were differentially regulated. These effects suggest that infection with C. parvum alters the host biochemical pathways by affecting gene expression (Deng et al., 2004).
V. A.
Human Infectivity Trials Prepatent and Patent Periods
Experimental data show that the parasite can complete its developmental cycle in 3 days (Current and Haynes, 1984). A study on the infectivity of C. parvum in healthy hosts revealed that oocysts can be detected in the stool as early as 2 days after ingestion of the inoculum, although most infections are detected 5 to 10 days after ingestion (prepatent period: 2 to 24 days) (Dupont et al., 1995). The duration of oocyst shedding ranged from 1 to 38 days. Of 18 volunteers who became infected, 11 developed enteric symptoms, 7 with diarrhea, thus meeting the study’s criteria for clinical cryptosporidiosis. The mean incubation period for diarrheal illness was 9 days (range: 4 to 22 days), and the mean duration of illness was 74 h (range: 58 to 87 h). Further analyses revealed that volunteers with diarrhea or enteric symptoms excreted more oocysts or were more likely to shed oocysts on consecutive days (Chappell et al., 1996). A recent study of C. hominis infection in healthy adults reported a trend toward a shorter incubation period (mean ± SD, 5.4 ± 2.7 days) and longer duration of illness (137 ± 142.3 h) but relatively less severe illness as measured by the number of unformed stools and the total unformed stool weight per diarrheal episode, compared to C. parvum infection (Table 8.2) (Dupont 1995; Chappell et al., 2006). An earlier study of naturally occurring human cryptosporidiosis revealed an incubation period of 7.2 days (range: 1 to 12 days) and a duration of illness of 12.2 days (range: 2 to 26 days) (Jokipii and Jokipii, 1986). In 19 of 26 patients, oocyst shedding persisted for a mean period of 6.9 days (range: 1 to 15 days) after cessation of symptoms. In 3 of 14 patients, oocyst excretion was detected up to 2 months after clinical recovery.
B.
Infectious Dose
Infection is initiated when the host ingests oocysts. Although the infectious dose for many animal species is not known, inoculation of 100 to 500 C. parvum oocysts caused infection in 50% of Swiss–Webster mice (Ernest et al., 1986). A report of a researcher who became infected after Cryptosporidium was coughed on him suggests that the infectious dose is low for humans (Blagburn and Current, 1983). Indeed, in an earlier study, the ID50 of C. parvum (Iowa isolate) was determined to be 132 oocysts among healthy volunteers (Dupont et al., 1995). Further investigation revealed heterogeneity in infectivity among
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TABLE 8.2 Clinical Features of Healthy Adults Infected with either C. parvum or C. hominis Oocysts
Clinical Features
C. parvum, 30–106/dose (29 adults)
C. hominis 10–500/dose (21 adults)
Infection rate Enteric symptoms rateb Diarrhea attack ratec Incubation period (mean days) Duration of illness (mean hours) Maximal number of unformed stools per day (mean) Number of unformed stools per illness (mean) Total stool weight per episode of diarrhea (mean, kg)
18(62%) 11(38%) 7(24%)d 9 74 6.4 12.7 1.23
9(43%) 14(67%) 13(62%)e 5.4 137.3 3.2 8.9 1.08
a
a
b
c
d e
Infection was defined as excretion of oocysts in stool more than 36 h after ingestion of oocysts. Rate of development of fever, nausea, vomiting, abdominal pain or cramps, tenesmus, gas-related intestinal symptoms, fecal urgency, or fecal incontinence. Diarrhea was defined as passage of 3 unformed stools in 8 h or of more than 3 unformed stools in 24 h in addition to the presence of at least one enteric symptom in the C. parvum study and passage of ≥ 200 g of unformed stool per day, ≥ 3 unformed stools in 8 h, or ≥ unformed stools in 24 h in the C. hominis study. All of these volunteers excreted oocysts. Of the 13 volunteers, 6 had detectable oocysts in the stool.
Source: Adapted from Dupont 1995 and Chappell 2006. With permission.
isolates of this species (Okhuysen et al., 1999). Three distinct C. parvum isolates had different ID50 levels—Iowa (calf), 87; UCP (calf), 1042; and TAMU (horse), 9. In the study of C. hominis infection of human volunteers, the ID50 was as low as 10 oocysts (range: 10 to 83) (Chappell et al., 2006). In contrast to C. parvum infection in human volunteers, C. hominis elicited a serum IgG response in most infected persons. From mathematical modeling of the Milwaukee outbreak, it was estimated that the infectious dose might have been as low as one oocyst for some persons (MacKenzie et al., 1994; Centers for Disease Control, 1995).
C.
Clinical versus Subclinical Infections
Until recently, asymptomatic carriage of the oocysts in clinically unaffected populations was thought to be infrequent. For example, in three studies examining predominantly asymptomatic homosexual men attending sexually transmitted disease clinics during the early 1980s, none of 375 individuals had Cryptosporidium oocysts in their stools. In contrast, Giardia lamblia was found in 3.3 to 6.5% and Entamoeba dispar was found in 5.5 to 23.5% of the men (McMillan and McNeillage, 1984; Jokipii et al., 1985; Chaisson et al., 1985). The prevalence of infection (other than outbreaks) with Cryptosporidium in developed countries such as the United States, Australia, or Europe average 2.1% in symptomatic individuals (range: 0.26 to 22%) and 0.2% in controls (Adal et al., 1995). In contrast, in developing countries, rates average 6.1% in symptomatic individuals (range:1.4 to 40.9%) and 1.5% in controls (Addy and Aikins-Bekoe, 1986; van den Ende, 1986; Pape et al., 1987; Guerrant, 1997). Persistent asymptomatic oocyst excretion can extend beyond clinical illness (Hart et al., 1984; Baxby et al., 1985; Ratnam et al., 1985; Jokipii and Jokipii, 1986; Stehr-Green et al., 1987). Asymptomatic cryptosporidiosis may be more common in the immunocompromised than in the immunocompetent hosts (PettoelloMantovani et al., 1995; Turkcapar et al., 2002; Houpt et al., 2005).
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Cryptosporidium and Cryptosporidiosis, Second Edition Postinfection Sequelae in Healthy Hosts
Musculoskeletal symptoms occurring within 1 to 2 weeks of cryptosporidial infection were described among immunocompetent hosts (Hay et al., 1987; Shepherd et al., 1988; Shepherd et al., 1989; Cron and Sherry, 1995; Ozgul et al., 1999; Collings and Highton, 2004). Reactive arthritis of the wrist, interphalangeal joints, knee, sacroiliac joint, or ankle occurred in both adult patients and children, most of whom were negative for HLA-B27. In a follow-up study of 111 cases of laboratory-confirmed cryptosporidiosis in England and Wales, symptoms of joint pain (odds ratio [OR], 2.8), eye pain (OR, 2.44), recurrent headache (OR, 2.1), dizzy spells (OR, 1.69), and fatigue (OR, 3.0) were more commonly reported by case patients than by age-matched control subjects who did not have gastrointestinal illness (Hunter et al., 2004). Interestingly, although 40% of case patients reported relapse of gastrointestinal symptoms after initial cryptosporidial illness irrespective of the genotype of the isolated organism, nongastrointestinal symptoms was noted only among those who had C. hominis and not C. parvum infection. A similar survey of self-reported complications of enteric infections from laboratory-confirmed cases reported to the Foodborne Diseases Active Surveillance Network (FoodNet) in California found that 8% of the 153 respondents with new symptoms after their gastrointestinal illness had joint pains (Rees et al., 2004). In contrast to the European study, none of the 35 patients who had Cryptosporidium (genotype not specified) and had completed the survey experienced rheumatic symptoms. Altered bowel habits, including persistent diarrhea of at least 3 months’ duration, were seen in 28.6% of the cases of cryptosporidial infection.
VI. Conclusion Cryptosporidium infection remains a debilitating illness among the immunocompromised hosts and very young children. Although the molecular mechanisms for the organism’s interaction with the host cell to cause acute disease are beginning to be clarified, more studies are needed to elucidate the mechanisms involved in the development of persistent diarrhea and long-term effects of infection in the growth and development of children. The epidemiologic and clinical features of cryptosporidial infection among the elderly, as well as the gastrointestinal and nongastrointestinal sequelae in immunocompetent hosts warrant further investigation. The relationship of the organism’s genotype with clinical disease is yet to be defined.
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Neill, M.A., Rice, S.K., Ahmad, N.V. and Flanigan, T.P. 1996. Cryptosporidiosis: An unrecognized cause of diarrhea in elderly hospitalized patients. Clin. Infect. Dis. 22, 168–170. Newman, R.D., Sears, C.L., Moore, S.R., Nataro, J.P., Wuhib, T., Agnew, D.A., Guerrant, R.L. and Lima, A.A. 1999. Longitudinal study of Cryptosporidium infection in children in northeastern Brazil. J. Infect. Dis. 180, 167–175. Niehaus, M.D., Moore, S.R., Patrick, P.D., Derr, L.L., Lorntz, B., Lima, A.A. and Guerrant, R.L. 2002. Early childhood diarrhea is associated with diminished cognitive function 4 to 7 years later in children in a northeast Brazilian shantytown. Am. J. Trop. Med. Hyg. 66, 590–593. Oberhuber, G., Laurer, E., Stolte, M. and Borchard, F. 1991. Cryptosporidiosis of the appendix vermiformis: A case report. Z. Gastroenterol. 29, 606–608. Ojcius, D.M., Perfettini, J.L., Bonnin, A. and Laurent, F. 1999. Caspase-dependent apoptosis during infection with Cryptosporidium parvum. Microbes Infect. 1, 1163–1168. Okhuysen, P.C., Chappell, C.L., Crabb, J.H., Sterling, C.R. and DuPont HL. 1999. Virulence of three distinct Cryptosporidium parvum isolates for healthy adults. J. Infect. Dis. 180, 1275–1281. Orenstein, J.M. 1997. Cryptosporidiosis, in D.H. Connor, F.W. Chandler, H.J. Manz, D.A. Schwartz and E.E. Lack (Eds.), Pathology of Infectious Diseases, Appleton & Lange, Stamford, CT, 1147–1158. Ozgul, A., Tanyuksel, M., Yazicioglu, K. and Arpacioglu, O. 1999. Sacroiliitis associated with Cryptosporidium parvum in an HLA-B27-negative patient. Rheumatology (Oxford). 38,288–289. Pape, J.W., Levine, E., Beaulieu, M.E., Marshall, F., Verdier, R. and Johnson, W.D. Jr. 1987. Cryptosporidiosis in Haitian children. Am. J. Trop. Med. Hyg. 36, 333–337. Pandak, N., Zeljka, K. and Cvitkovic A. 2006. A family outbreak of cryptosporidiosis: Probable nosocomial infection and person-to-person transmission. Wien. Klin. Wochenschr. 118, 485–487. Perch, M., Sodemann, M., Jakobsen, M.S., Valentiner-Branth, P., Steinsland, H., Fischer, T.K., Lopes, D.D., Aaby, P. and Molbak, K. 2001. Seven years' experience with Cryptosporidium parvum in Guinea-Bissau, West Africa. Ann. Trop. Paediatr. 21,313–318. Pereira, M.D., Atwill, E.R., Barbosa, A.P., Silva, S.A., and Garcia-Zapata, M.T. 2002. 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. Petras, R.E., Carey, W.D. and Alanis, A. 1983. Cryptosporidial enteritis in a homosexual male with an acquired immunodeficiency syndrome. Cleve. Clin. Q. 50, 41–45. Pettoello-Mantovani, M., Di Martino, L., Dettori, G., Vajro, P., Scotti, S., Ditullio, M.T. and Guandalini, S. 1995. Asymptomatic carriage of intestinal Cryptosporidium in immunocompetent and immunodeficient children: A prospective study. Pediatr Infect Dis J. 14,1042–1047. Priest, J.W., Li, A., Khan, M., Arrowood, M.J., Lammie, P.J., Ong, C.S., Roberts, J.M. and Isaac-Renton, J. 2001. Enzyme immunoassay detection of antigen-specific immunoglobulin g antibodies in longitudinal serum samples from patients with cryptosporidiosis. Clin. Diagn. Lab. Immunol. 8, 415–423. Ratnam, S., Paddock, J., McDonald, E., Whitty, D., Jong, M. and Cooper, R. 1985. Occurrence of Cryptosporidium oocysts in fecal samples submitted for routine microbiological examination. J. Clin. Microbiol. 22, 402–404. Rees, J.R., Pannier, M.A., McNees, A., Shallow, S., Angulo, F.J. and Vugia, D.J. Persistent diarrhea, arthritis, and other complications of enteric infectiojns: A pilot survey based on California FoodNet surveillance, 1998–1999. 2004. Clin. Infect. Dis. 38, S311–317. Ribeiro, C.D. and Palmer, S.R.1986. Family outbreak of cryptosporidiosis. Br. Med. J. 292, 377. Robinson, P., Okhuysen, P.C., Chappell, C.L., Lewis, D.E., Shahab, I., Janecki, A. and White, A.C. Jr. 2001. Expression of tumor necrosis factor alpha and interleukin 1 beta in jejuna of volunteers after experimental challenge with Cryptosporidium parvum correlates with exposure but not with symptoms. Infect Immun. 69,1172–1174. Robinson, P., Okhuysen, P.C., Chappell, C.L., Weinstock, J.V., Lewis, D.E., Actor, J.K. and White, A.C. Jr. 2003. Substance P expression correlates with severity of diarrhea in cryptosporidiosis. J. Infect. Dis. 188, 290–296. Epub 2003 July 9. Rocha, F., Musch, M.W., Lishanskiy, L., Bookstein, C., Sugi, K., Xie, Y. and Chang, E.B. 2001. IFN-gamma downregulates expression of Na(+)/H(+) exchangers NHE2 and NHE3 in rat intestine and human Caco2/bbe cells. Am. J. Physiol. Cell. Physiol. 280, C1224–C1232.
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Rodriguez-Hernandez, J., Canut-Blasco, A. and Martin-Sanchez, A.M. 1996. Seasonal prevalences of Cryptosporidium and Giardia infections in children attending day care centres in Salamanca (Spain) studied for a period of 15 months. Eur. J. Epidemiol. 12, 291–295. Rossi, P., Rivasi, F., Codeluppi, M., Catania, A., Tamburrini, A., Righi, E. and Pozio, E. 1998. Gastric involvement in AIDS associated cryptosporidiosis. Gut. 43, 476–477. Sallon, S., Deckelbaum, R.J., Schmid, I.I., Harlap, S., Baras, M. and Spira, D.T. 1988. Cryptosporidium, malnutrition, and chronic diarrhea in children. Am. J. Dis. Child. 142, 312–315. Sallon, S., el-Shawwa, R., Khalil, M., Ginsburg, G., el Tayib, J., el-Eila, J., Green, V. and Hart, C.A. 1994. Diarrhoeal disease in children in Gaza. Ann. Trop. Med. Parasitol. 88,175–182. Samson, V.E. and Brown, W.R. 1996. Pneumatosis cystoides intestinalis in AIDS-associated cryptosporidiosis. More than an incidental finding? J. Clin. Gastroenterol. 22, 311–312. Sarabia-Arce, S., Salazar-Lindo, E., Gilman, R.H., Naranjo, J. and Miranda, E. 1990. Case-control study of Cryptosporidium parvum infection in Peruvian children hospitalized for diarrhea: Possible association with malnutrition and nosocomial infection. Pediatr. Infect. Dis. J. 9, 627–631. Schneiderman, D.J., Cello, J.P. and Laing, F.C. 1987. Papillary stenosis and sclerosing cholangitis in the acquired immunodeficiency syndrome. Ann. Intern. Med. 106,546–549. Schoofs, M.W., Maartense, E., Eulderink, F., Vreede, R.W. 2004. Cryptosporidiosis leading to an unsuspected diagnosis of AIDS. Neth. J. Med. 62,198–200. Sears, C.L. and Guerrant, R.L. 1994. Cryptosporidiosis: The complexity of intestinal pathophysiology. Gastroenterology. 106, 252–254. Seydel, K.B., Zhang, T., Champion, G.A., Fichtenbaum, C., Swanson, P.E., Tzipori, S., Griffiths, J.K. and Stanley, S.L. Jr. 1998. Cryptosporidium parvum infection of human intestinal xenografts in SCID mice induces production of human tumor necrosis factor alpha and interleukin-8. Infect. Immun. 66, 2379–2382. Shepherd, R.C., Sinha, G.P., Reed, C.L. and Russell, F.E. 1988. Cryptosporidiosis in the West of Scotland. Scot. Med. J. 33,365–368. Shepherd, R.C., Smail, P.J. and Sinha, G.P. 1989. Reactive arthritis complicating cryptosporidial infection. Arch. Dis. Child. 64, 743–744. Sidhu, S., Flamm, S. and Chopra, S. 1994. Pneumatosis cystoides intestinalis: An incidental finding in a patient with AIDS and cryptosporidial diarrhea. Am. J. Gastroenterol. 89, 1578–1579. Smith, H.V., Nichols, R.A. and Grimason, A.M. 2005. Cryptosporidium excystation and invasion: Getting to the guts of the matter. Trends Parasitol. 21, 133–142. Soave, R., Danner, R.L., Honig, C.L., Ma, P., Hart, C.C., Nash, T. and Roberts, R.B. 1984. Cryptosporidiosis in homosexual men. Ann. Intern. Med. 100, 504–511. Soave, R. and Ma, P. 1985. Cryptosporidiosis: Traveler’s diarrhea in two families. Arch. Intern. Med. 145:70–72. Soave, R. and Armstrong, D. 1986. Cryptosporidium and cryptosporidiosis. Rev. Infect. Dis. 8, 1012–1023. Review. Erratum in: Rev. Infect. Dis. 1987 May-Jun, 9(3):663. de Souza, Ldo. R., Rodrigues, M.A., Morceli, J., Kemp, R. and Mendes, R.P. 2004. Cryptosporidiosis of the biliary tract mimicking pancreatic cancer in an AIDS patient. Rev. Soc. Bras. Med. Trop. 37,182–185. Epub 2004 April 13. Sreedharan, A., Jayschree, R.S. and Sridhar, H. 1996. Cryptosporidiosis among cancer patients: An observation. J. Diarrhoeal Dis. Res. 14,211–213. Stehr-Green, J.K., McCaig, L., Remsen, H.M., Rains, C.S., Fox, M. and Juranek DD. 1987. Shedding of oocysts in immunocompetent individuals infected with Cryptosporidium. Am. J. Trop. Med. Hyg. 36, 338–342. Sterling, C.R., Gilman, R.H., Sinclair, N.A., Cama, V., Castillo, R. and Diaz, F. 1991. The role of breast milk in protecting urban Peruvian children against cryptosporidiosis. J. Protozool. 38, 23S–25S. Sugi, K., Musch, M.W., Field, M. and Chang, E.B. 2001. Inhibition of Na+, K+-ATPase by interferon gamma down-regulates intestinal epithelial transport and barrier function. Gastroenterology. 120, 1393–1403. Tanyuksel, M., Gun, H. and Doganci, L. 1995. Prevalence of Cryptosporidium sp. in patients with neoplasia and diarrhea. Scand. J. Infect. Dis. 27, 69–70. Teixidor, H.S., Godwin, T.A. and Ramirez, E.A. 1991. Cryptosporidiosis of the biliary tract in AIDS. Radiology. 180, 51–6. Teresa Ortega, M., Vergara, A., Guimbao, J., Clavel, A., Gavin, P. and Ruiz, A. 2006. Cryptosporidium hominis diarrhea outbreak and transmission linked to diaper infant use. Med. Clin. (Barc). 127, 653–656.
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Topouchian, A., Kapel, N., Huneau, J.F., Barbot, L., Magne, D., Tome, D. and Gobert, JG. 2001. Impairment of amino-acid absorption in suckling rats infected with Cryptosporidium parvum. Parasitol Res. 87, 891–896. Tumwine, J.K., Kekitiinwa, A., Bakeera-Kitaka, S., Ndeezi, G., Downing, R., Feng, X., Akiyoshi, D.E. and Tzipori, S. 2005. Cryptosporidiosis and microsporidiosis in Ugandan children with persistent diarrhea with and without concurrent infection with the human immunodeficiency virus. Am. J. Trop. Med. Hyg. 73, 921–925. Tumwine, J.K., Kekitiinwa, A., Nabukeera, N., Akiyoshi, D.E., Rich, S.M., Widmer, G., Feng, X. and Tzipori, S. Cryptosporidium parvum in children with diarrhea in Mulago Hospital, Kampala, Uganda. Am. J. Trop. Med. Hyg. 2003. 68, 710–715. Turkcapar, N., Kutlay, S., Nergizoglu, G., Atli, T. and Duman, N. 2002. Prevalence of Cryptosporidium infection in hemodialysis patients. Nephron. 90, 344–346. Tzipori, S. 1983. Cryptosporidiosis in animals and humans. Microbiol. Rev. 47, 84–96. Tzipori, S., Angus, K.W., Campbell, I and Gray, E.W. 1980. Cryptosporidium: Evidence for a single-species genus. Infect. Immun. 30, 884–886. Vakil, N.B., Schwartz, S.M., Buggy, B.P., Brummit, C.F., Kherellah, M., Letzer, D.M., Gilson, I.H. and Jones, P.G. 1996. Biliary cryptosporidiosis in HIV-infected people after the waterborne outbreak of cryptosporidiosis in Milwaukee. N. Engl. J. Med. 334,19–23. van den Ende, GM .1986. Cryptosporidiosis among black children in hospital in South Africa. J. Infect. Dis. 13, 25–30. Ventura, G., Cauda, R., Larocca, M., Riccioni, M.E., Tumbarello, M. and Lucia, M.B. 1997. Gastric cryptosporidiosis complicating HIV infection: Case report and review of the literature. Eur. J. Gastroenterol. Hepatol. 9, 307–310. Vetterling, J.M., Takeuchi, A. and Madden, P.A. 1971. Ultrastructure of Cryptosporidium wrairi from the guinea pig. J. Protozool. 18, 248–260. Westrope, C. and Acharya, A. 2001. Diarrhea and gallbladder hydrops in an immunocompetent child with Cryptosporidium infection. Pediatr. Infect. Dis. J. 20, 1179–1181. Wolfson, J.S., Richter, J.M., Waldron, M.A., Weber, D.J., McCarthy, D.M. and Hopkins, C.C. 1985. Cryptosporidiosis in immunocompetent patients. N. Engl. J. Med. 312, 1278–1282. Zhang, Y., Lee, B., Thompson, M., Glass, R., Cama, R.I., Figueroa, D., Gilman, R., Taylor, D. and Stephenson, C. 2000. Lactulose-mannitol intestinal permeability test in children with diarrhea caused by rotavirus and cryptosporidium. Diarrhea Working Group, Peru. J. Pediatr. Gastroenterol. Nutr. 31, 16–21. Erratum in: J. Pediatr. Gastroenterol. Nutr. 2000 Nov; 31(5):578. Zu, S.X., Li, J.F., Barrett, L.J., Fayer, R., Shu, S.Y., McAuliffe, J.F., Roche, J.K. and Guerrant, R.L. 1994. Seroepidemiologic study of Cryptosporidium infection in children from rural communities of Anhui, China and Fortaleza, Brazil. Am. J. Trop. Med. Hyg. 51, 1–10.
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9 Prophylaxis and Chemotherapy
Heather D. Stockdale, Jennifer A. Spencer, and Byron L. Blagburn
CONTENTS I. II.
Introduction .................................................................................................................................. 255 Prophylaxis and Chemotherapy in Humans ................................................................................ 256 A. Clinical Manifestations ................................................................................................... 256 B. Chemotherapeutic Agents ............................................................................................... 256 1. Macrolides........................................................................................................ 257 2. Rifabutin........................................................................................................... 261 3. Benzeneacetonitrile Derivatives....................................................................... 261 4. Miscellaneous Agents ...................................................................................... 262 C. Highly Active Antiretroviral Therapy (HAART) ........................................................... 263 D. Supportive Therapy ......................................................................................................... 264 E. Future Directions............................................................................................................. 265 III. Treatment and Prophylaxis in Animals........................................................................................ 265 References........................................................................................................................... 279
I.
Introduction
We have experienced enormous advances in our knowledge of the biology, pathogenesis, diagnosis, and control of human and animal cryptosporidiosis during the decade preceding the first printing of this book (Thompson et al., 2005). Indeed, one could presume that much of what was once speculative regarding Cryptosporidium is now better understood. Certainly, research has helped us to understand the complex taxonomic interrelationships between the different species and genotypes of Cryptosporidium. We also now enjoy improved diagnostic tests and also have a better understanding of the epidemiology of human and animal disease caused by Cryptosporidium spp. (Sunnotel et al., 2006). However, chemotherapy of cryptosporidiosis is one area of research in which advances have been less dramatic. The reasons for the slower progress are complex and relate to the uniqueness of Cryptosporidium among apicomplexan protists and the appearance of highly effective viral therapies for HIV infection and AIDS (Cacciò and Pozio, 2006). Regarding the former, some researchers now consider Cryptosporidium more closely related to the Gregarinidae than the classical coccidia, based on the biological behavior of Cryptosporidium in cell-free culture systems (Hijjawi et al., 2004) and continuing phylogenetic analysis (Morrison et al., 2004). The extent to which their atypical responses to traditional anticoccidial medications can be attributed to their unique taxonomic placement is yet to be determined. In this chapter, as with the previous edition, we have elected to survey and compile the available scientific literature regarding Cryptosporidium chemotherapy. We include both successful and unsuccessful attempts to inactivate Cryptosporidium using both chemical and immunologic intervention strategies. We have retained much of the older literature summarized in the previous iteration of this chapter (Blagburn and Soave, 1997). We do so because we feel that a comprehensive compilation of available information would be helpful to those new to the field. We have also retained the table format because we feel that 255
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it is the most efficient method of dealing with the volume of literature in the field. We have also added a table to summarize relevant human therapies (missing in the earlier edition) and a section on highly active antiretroviral therapy (HAART).
II. A.
Prophylaxis and Chemotherapy in Humans Clinical Manifestations
Human clinical cryptosporidial infection ranges from mild, self-limited diarrhea to fulminant, choleralike enteritis complicated by extraintestinal disease (see also Chapter 8). Asymptomatic infection occurs, but the frequency is unknown. Clinical manifestations of symptomatic cryptosporidial infection include watery diarrhea, abdominal pain, nausea, vomiting, flatulence, bloating, urgency, incontinence, anorexia, and weight loss. The severity and duration of human cryptosporidial disease varies with host immune competence (Soave et al., 1984; Wolfson et al., 1985; Chapter 7). When immune function is intact, cryptosporidiosis is usually explosive at onset, lasts approximately 10 to 14 days, and is followed by a complete clinical and parasitologic recovery. Oocyst shedding may lag behind clinical resolution of symptoms by as long as a few weeks. In the immunologically impaired host, onset of cryptosporidiosis is usually insidious; diarrheal symptoms precede detection of the organism in stool by weeks to months. In these individuals, the enteritis becomes chronic, and severity may wax and wane. In persons with the acquired immunodeficiency syndrome (AIDS), diarrhea frequency and volume often escalate unrelentingly as immune function becomes more deranged. However, cryptosporidiosis may also resolve spontaneously in patients anywhere along the spectrum of HIV infection, thus complicating the interpretation of uncontrolled treatment data. For the most part, cryptosporidiosis is a devastating complication for persons with AIDS; it contributes significantly to morbidity and hastens death (Mannheimer and Soave, 1994; Petersen, 1992). In addition to AIDS, immunologic deficiencies and other conditions associated with protracted cryptosporidiosis include congenital hypogammaglobulinemia, IgA deficiency, concurrent viral infections, malnutrition, and exogenous immunosuppression, as with corticosteroids (Current and Garcia, 1991; Ungar, 1994). If exogenous immunosuppression can be discontinued, cryptosporidiosis will resolve. Biliary cryptosporidiosis has been well documented only in the immunocompromised host. Because definitive diagnosis requires invasive procedures that may not be justified in the absence of useful treatment options, the true incidence of this complication is not known (Bouche et al., 1993; Gross et al., 1986; Hinnant et al., 1989; Teixidor et al., 1991). Pancreatitis and reactive arthritis associated with cryptosporidiosis have also been described in immunocompetent patients as well as those with AIDS (Gross et al., 1986; Hinnant et al., 1989; Miller et al., 1992; Shepherd et al., 1989). Respiratory cryptosporidiosis, well recognized in birds has been reported in a few humans, but in these cases, contamination of respiratory secretions from intestinal infection has not been satisfactorily ruled out (Current and Garcia, 1991; Fayer and Ungar, 1986; Mannheimer and Soave, 1994; Ungar, 1994).
B.
Chemotherapeutic Agents
There have been great strides in improving the treatment of cryptosporidiosis in humans over the past decade. In addition to advances in effective therapy, the availability and reproducibility of in vivo and in vitro methods for screening drugs and conducting preclinical studies has greatly enhanced efforts to identify effective therapy. Progress has been made in developing animal models of the disease, but still little is known of how cryptosporidial species and strain differences impact parasite virulence. Despite the paucity of supportive preclinical data, the urgent need to identify effective therapy for this disease in persons with AIDS has led to the unprecedented administration of a vast array of chemotherapeutic, immunomodulatory, and palliative agents to this population. The largely anecdotal experience thus generated has resulted in a formidable list of approximately 100 ineffective compounds (Fayer and Ungar, 1986; Soave, 1990). Over the past two decades, controlled treatment trials have provided useful
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insights for designing effective studies of cryptosporidial therapy, and there is now an approved, effective treatment for cryptosporidiosis in humans. The following is a compilation of chemotherapeutic and palliative attempts at treating cryptosporidiosis (Table 9.1). Immunomodulatory therapy has been covered in Chapter 7.
1. a.
Macrolides Oral Spiramycin (Rovamycine®, Rovamicina®, Provamicina®, Rhône–Poulenc Rorer)
Initially, interest centered on spiramycin, a macrolide discovered in 1957 and, though not licensed in the United States, used worldwide. Spiramycin has broad spectrum antibacterial activity but interest in its potential as an anticryptosporidial agent derives from its activity against a related protozoan, Toxoplasma gondii. In the mid-to-late 1980s, there were at least five anecdotal reports of success and failure with spiramycin for cryptosporidiosis (Anonymous, 1984; Collier et al., 1984; Fafard and Lalonde, 1990; Moskovitz et al., 1988; Portnoy et al., 1984). These divergent results led to the first controlled cryptosporidiosis treatment trials, but they, too, provided divergent data. In a prospective study, 15 patients were randomized to either spiramycin or erythromycin; both antibiotics were associated with clinical response, but side effects limited their use (Connolly et al., 1988). In a placebo-controlled “n of 1” trial, no difference was shown between spiramycin and placebo (Woolf et al., 1987). There have been two placebo-controlled trials of spiramycin for cryptosporidiosis in immunocompetent children, and one in adults with AIDS. In 39 immunocompetent infants and children in Durban, South Africa, randomized to 5 days of treatment with either spiramycin (75 mg/kg) or placebo (Wittenberg et al., 1989), no difference was found between active drug and placebo, possibly because the dose was too low or the duration of therapy was too short. In a study of 44 immunocompetent infants in Costa Rica randomized to spiramycin (100 mg/kg/day) or matching placebo (which contained lactose) for 10 days, a statistically significant decrease in diarrhea and oocyst excretion was found in the spiramycin-treated group (Saez-Llorens et al., 1989). In a randomized, double-blind, placebo-controlled trial, 3.0 MIU (approximately 3 g) spiramycin thrice daily for 3 weeks was no better than the placebo in the treatment of cryptosporidial diarrhea in 73 patients with AIDS (Blagburn and Soave, 1997). A food interaction study in normal hosts indicated that spiramycin absorption was significantly decreased in the presence of food, perhaps explaining the poor results obtained in the controlled trial with AIDS patients. Interestingly, less rigorously obtained data from the open-label compassionate use program that ran concomitantly with the placebo-controlled study revealed that 28 of 37 AIDS patients had a favorable clinical response and 12 had parasitologic eradication, thus underscoring the importance of placebo-controlled clinical treatment trials (Moskovitz et al., 1988).
b.
Intravenous Spiramycin
To circumvent absorption problems, an efficacy and safety study of intravenous spiramycin was conducted in 1989–91. In this multicenter, single-blind, placebo-controlled, National Institutes of Allergy and Infectious Diseases (NIAID)-sponsored AIDS Clinical Trials Group (ACTG #113) trial, 2 doses of intravenous spiramycin (3.0 MIU and 4.5 MIU) were examined. Of 31 AIDS patients enrolled, 5 (18%) had a favorable response to treatment, i.e., both clinical and parasitologic improvement, whereas 16 (57%) had partial benefit but did not meet the favorable response criteria. Analysis of the group revealed a statistically significant drop in oocyst numbers while receiving spiramycin as compared to placebo (Blagburn and Soave, 1997). However, administration of intravenous spiramycin was associated with paresthesias, taste perversion, nausea, and vomiting, and at doses greater than 75 mg/kg/day, there were cases of severe colitis due to intestinal injury (Weikel et al., 1991).
c.
Azithromycin (Zithromax®, Pfizer)
Azithromycin is an azalide with a long half-life (6–8 h) that achieves high tissue concentrations, particularly in the biliary tree and gallbladder. Experience with azithromycin for cryptosporidiosis in humans includes anecdotal data, prospective and placebo-controlled studies, and open-label protocols. According to a 1996 prospective study, azithromycin does not seem to have a prophylactic effect against
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TABLE 9.1 Efficacies of Potential Anticryptosporidial and Antiretroviral Therapies (Separately or in Combination) in Humans Age Adult
Adult Adult Adult
Adult
Adult
Health Status
Drug
HIV; CD4+ counts Clarithromycin < 0.75 × 109/L Rifabutin Azithromycin AIDS; CD4+ Clarithromycin counts < 50/mm3 HIV; CD4+ counts Clarithromycin < 100 × 106/L Rifabutin CD30+ anaplastic Paromomycin large-cell lymphoma Nodular sclerosing rh IL-12 Hodgkin’s disease Paromomycin Azithromycin AIDS; CD4+ Azithromycin counts < 100/µL Letrazuril Paromomycin
Adult Adult
HIV; CD4+ counts Paromomycin < 150/mm3 AIDS; CD4+ Azithromycin counts 0–85 × 106/L, one 162 × 106/L
Adult
HIV
Azithromycin
Adult
HIV; CD4+ counts 120/mm3 Acute lymphoblastic leukemia
Azithromycin Paromomycin Paromomycin
Adult
AIDS; CD4+ counts < 100/µL (8–55/mm3)
Paromomycin
Child
Normal
Azithromycin
Child
Azithromycin
Praziquantel Mirazid
Method of Treatment Prophylactic or and Dosage Efficacy Therapeutic Use
Reference
Not specified
+/–
Prophylactic
Holmberg et al., 1998
500 mg BID
+
Prophylactic
Jordan, 1996
500 mg BID 300–450 mg daily 4 g/day
+/–
Prophylactic
–
Therapeutic
Fichtenbaum et al., 2000 Nachbaur et al., 1997
SQ 4.5 × 106 I U/day for 4 days, repeated 3 weeks later 4 g/day orally 250 mg BID 1 g or 500 mg for 2–4 weeks 50–200 mg up to 4 weeks 250–500 mg QID for 1 month 500 mg QID for 21 days 500 mg/day for 30–60 days 1000 mg/day for 20–50 days 1500 mg/day for 20 days 500 mg/day for 5–14 days 500 mg BID orally 500 mg BID orally 35 mg/kg/day for 10 days 20 mg/kg/day for 14 days Treatments were given alone or in combination 1.0 g BID for 4 or 8 weeks 600 mg PID for 4 weeks 500 mg/day for 3 days orally 40 mg/kg single dose orally 10 mg/kg/day for 3 days orally
+/–
Therapeutic
Nachbaur et al., 1997
+/–
Therapeutic
Blanshard et al., 1997
–
Therapeutic
+/–
Therapeutic
Hewitt et al., 2000 Dionisio et al., 1998
+/–
Therapeutic
+
Therapeutic
+
Therapeutic
+
Therapeutic
Smith et al., 1998
+/–
Therapeutic
Allam and Shehab, 2002
Kadappu et al., 2002 Meamar et al., 2006 Trad et al., 2003
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TABLE 9.1 (CONTINUED) Efficacies of Potential Anticryptosporidial and Antiretroviral Therapies (Separately or in Combination) in Humans Age
Health Status
Adult/ AIDS adolescent Adult HIV
Drug Nitazoxanide Nitazoxanide
Adult/ Normal adolescent Child Normal
Nitazoxanide
Adult
AIDS
Roxithromycin
Adult
HIV
Roxithromycin
Adolescent
Coeliac disease
Adult
Adult
Adult
Nitazoxanide
Lactobacillus GG Lactobacillus casei Shirota AIDS Glycine Glutamine + glycine HIV; CD4+ counts Glutamine 40/mm3 Azithromycin Paromomycin HAART Stavudine Lamivudine Indinavir HIV; CD4+ count Paromomycin 45/µL HAART Zidovudine Zalcitabine Indinavir
Adult Adult
HIV; CD4+ count < 50 × 106/L HIV; CD4+ count 4–38 × 106/L
Indinavir HAART Stavudine Lamivudine Indinavir Zidovudine Saquinavir
Method of Treatment Prophylactic or and Dosage Efficacy Therapeutic Use 500 mg BID for 1 week orally 1 or 2 g/day
Reference
+
Therapeutic
+
Therapeutic
+
Therapeutic
+
Therapeutic
+
Therapeutic
+
Therapeutic
+
Therapeutic
Pickerd and Tuthill, 2004
46 g/day orally 30 g/day + 15 g/day orally 14 g daily added to parenteral nutrition 500 mg PID 500 mg TID
+/–
Therapeutic
Bushen et al., 2004
+/–
Therapeutic
Moling et al., 2005
250 mg QID for 9 days
+/–
Therapeutic
Schmidt et al., 2001
+
Therapeutic
+
Therapeutic
Foudraine et al., 1998 Miao et al., 1999
500 mg BID for 3 days given with food 5 mL (100 mg/5 mL) BID for 3 days with food 10 mL (100 mg/5 mL) BID for 3 days with food 300 mg BID for 4 weeks 300 mg BID for 4 weeks 109 units/day
Doumbo et al., 1997 Rossignol et al., 1998 Rossignol et al., 2001 Rossignol et al., 2001
Sprinz et al., 1998 Uip et al., 1998
6.5 × 109 units/day
250 mg BID for 4 weeks 0.75 mg TID for 4 weeks 800 mg TID for 4 weeks 2400 mg daily for 24 weeks Given as triple or quadruple therapy for 1–6 months
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TABLE 9.1 (CONTINUED) Efficacies of Potential Anticryptosporidial and Antiretroviral Therapies (Separately or in Combination) in Humans Age
Health Status
Adult
HIV-1
Adult
HIV-1; CD4+ count 7–179/µL
Drug HAART
Method of Treatment Prophylactic or and Dosage Efficacy Therapeutic Use
Given as double or triple therapy for 12 weeks Ritonavir 600 mg BID Saquinavir 400 mg BID or 600 mg TID Stavudine 40 mg BID Indinavir 800 mg TID Lamivudine 150 mg BID Paromomycin 500 mg QID for 1 Spiramycin month Azithromycin 6 × 106U QID for 1 HAART month Azidothymidine 500 mg TID for 1 Zalcitabine month Stavudine antiretrovirals given as Indinavir single, double, or Ritonavir triple therapies at Saquinavir standard dosages Lamivudine
Reference
+
Therapeutic
Carr et al., 1998
+/–
Therapeutic
Maggi et al., 2000
Note: + = Demonstrable activity against Cryptosporidium parvum; – = no demonstrable activity against C. parvum; +/– = partial activity at one or more dosages for a compound, or activity for one or more compounds in a group.
cryptosporidiosis in HIV patients (Holmberg et al., 1998). There have been several studies demonstrating the therapeutic effects of azithromycin. In a multicenter, double-blind, crossover trial, 85 AIDS patients with cryptosporidial diarrhea were randomized to receive either 900-mg azithromycin, once daily for 3 weeks, or matching placebo (Soave et al., 1993). Preliminary analysis of the data reveals no significant trend toward improvement in bowel movement frequency, stool oocyst shedding, and weight stabilization in the azithromycin-treated group. Data from the first 33 patients revealed a highly significant decrease in stool oocyst counts on days 7 and 21 in those subjects with the highest serum azithromycin levels (p < .01). A long-term study using azithromycin at low doses to treat HIV patients with chronic cryptosporidiosis demonstrated that the drug was well tolerated and could possibly eradicate the pathogen (Dionisio et al., 1998). Thirteen HIV patients with cryptosporidiosis were treated with 500, 1000, or 1500 mg of azithromycin over a period of 30 to 360 days, which included an initial treatment period and long-term maintenance period. Adverse reactions were seen only in patients given 1500 mg daily. Those patients taking 500 or 1000 mg showed no side effects and demonstrated a complete response during the initial treatment and when given 500 mg during the maintenance period up to 360 days (Dionisio et al., 1998). Using azithromycin to treat children with cryptosporidiosis also proved to be effective (Allam and Shehab, 2002). Eleven children were given 500 mg of azithromycin daily for three weeks resulting in a 91% cure rate and a 99% reduction in oocysts collected in stool samples. The complete resolution of cryptosporidial diarrhea was observed in two children, immunocompromised after receiving chemotherapy, treated with 40 mg/kg azithromycin (Vargas et al., 1993). In another study, children undergoing chemotherapy for acute lymphoblastic leukemia developed cryptosporidiosis and were successfully treated with azithromycin (Trad et al., 2003). These children were given 20 mg/kg per day for 14 days, which resolved diarrhea; stools were negative for Cryptosporidium and after 6–14 months showed no signs of relapse. To further investigate the effect of drug absorption in treatment of this disease, five very ill patients with AIDS and cryptosporidial diarrhea who had failed oral azithromycin were given intravenous azithromycin in a prospective, open-labeled, case-control study (Blagburn and Soave, 1997). Although
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bowel movement frequency and stool oocyst counts remained unchanged, serum alkaline phosphatase levels decreased in all 5 subjects who received intravenous azithromycin compared to 4 of 13 controls (p < 0.05). The data suggest a therapeutic effect on biliary tree infection but may be confounded by concomitant opportunistic infection with Mycobacterium avium complex or other organisms susceptible to azithromycin. Intravenous azithromycin at doses were well tolerated at up to 2 g daily for as long as 2 weeks.
d.
Clarithromycin (Biaxin®, Biaxin XL®, Abbott Laboratories) and Roxithromycin (Surlid®, Rulide®, Biaxsig®, Roxar®, Roximycin®, Hoechst Marion Roussel Ltd.)
Clarithromycin and roxithromycin are both semisynthetic macrolides. Clarithromycin, as with most macrolides, has a moderate bioavailability (55%), whereas roxithromycin has enhanced bioavailability (72–85%) (Zhanel et al., 2001). They have been used in clinical trials to determine their prophylactic and therapeutic effectiveness against cryptosporidiosis. Studies have indicated that clarithromycin has some prophylactic effect against cryptosporidiosis in immunocompromised patients. In a prospective study involving 136 AIDS patients, 63 received 500 mg of clarithromycin twice a day, and 73 did not receive clarithromycin. Those taking clarithromycin did not develop cryptosporidiosis, whereas 4 of 73 not taking clarithromycin did develop cryptosporidiosis (Jordan, 1996). There was also a subsequent study involving 217 AIDS patients taking clarithromycin for its prophylactic effects, and after a 2-year follow-up, no patient developed cryptosporidiosis (Jordan, 1996). Another prospective study indicates similar data regarding the prophylactic effectiveness of clarithromycin. There were 312 individuals with HIV given clarithromycin and only 5 developed cryptosporidiosis, whereas 30 of 707 HIV patients not taking clarithromycin developed cryptosporidiosis (Holmberg et al., 1998). One study reported conflicting results with clarithromycin as a prophylactic treatment. The data suggest that clarithromycin has no prophylactic effect against cryptosporidiosis in patients with HIV (Fichtenbaum et al., 2000). The use of roxithromycin as a therapeutic treatment for cryptosporidiosis in immunocompromised patients has proved effective. In an open study of 24 AIDS patients with cryptosporidiosis, roxithromycin was well tolerated with only minor side effects. A partial or complete response was seen in 79% of the patients, and those with a complete response remained negative for Cryptosporidium oocysts in stool samples during a 6-month follow-up (Sprinz et al., 1998). Similar results were found in another study in which 22 AIDS patients with cryptosporidiosis were treated with 600 mg of roxithromycin daily for 4 weeks. In this study, 68% had a complete response with 27% having an improved response (Uip et al., 1998).
2.
Rifabutin (Mycobutin®, Pfizer)
Rifabutin is a semisynthetic derivative of rifamycin S and well absorbed in all tissues (Farr, 2000; Kamps and Hoffman, 2006). Studies have shown that rifabutin given to HIV patients may help prevent the development of cryptosporidiosis. In a prospective study, 214 HIV patients were given rifabutin and only 2 became infected with Cryptosporidium over a 4-year period. There were also 805 HIV patients not taking rifabutin, and 33 developed cryptosporidiosis (Holmberg et al., 1998). Another study involving 650 HIV patients who were given 300 to 450 mg of rifabutin daily demonstrated a lower risk of developing cryptosporidiosis than if given clarithromycin alone or in combination with rifabutin (Fichtenbaum et al., 2000).
3. a.
Benzeneacetonitrile Derivatives Diclazuril (Clinacox®, Janssen Pharmaceuticals)
Interest in this class of agents for treatment of cryptosporidiosis stems from their activity against the related coccidian Eimeria. In addition to two anecdotal reports (Connolly et al., 1990; Menichetti et al., 1991), diclazuril sodium was tested in a randomized, double-blind, placebo-controlled, escalating dose trial in AIDS patients at three centers in New York City (Soave et al., 1990). Doses ranging from 50 to 800 mg were no more efficacious than placebo. However, the very small numbers of complete responders
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were the only subjects who had detectable serum diclazuril levels, suggesting that lack of efficacy may have been due to poor drug bioavailability.
b.
Letrazuril (Janssen Research Foundation)
Poor absorption of diclazuril led to synthesis of the p-fluor analog letrazuril. This agent was studied in a randomized, double-blind, placebo-controlled multicenter treatment trial with a pharmacokinetic arm (NIAID-ACTG 198). Letrazuril bioavailability was better than that of diclazuril, but patients experienced little clinical improvement. There was a highly significant parasitologic effect when stool specimens were examined by acid-fast staining, but specimens that were acid-fast negative were positive by ELISA antigen capture and indirect immunofluorescent assay, suggesting that letrazuril may interfere with acidfast staining of oocysts (Blagburn and Soave, 1997). Although data obtained in other open-label studies and anecdotal cases was more promising, in the majority, acid-fast staining of stool smears was used to assess drug parasitologic effects (Guillem et al., 1992; Hamour et al., 1993; Harris et al., 1994; Loeb et al., 1995; Murdoch et al., 1993; Victor et al., 1993). A pilot study examining the efficacy and safety of letrazuril was conducted involving AIDS patients with chronic cryptosporidiosis. Of the ten patients given 150 to 200 mg of letrazuril daily, four demonstrated a complete or partial response (Blanshard et al., 1997). The benzeneacetonitrile derivatives are not currently available for use in human cryptosporidiosis.
4. a.
Miscellaneous Agents Paromomycin (Humatin®, Parke-Davis)
Paromomycin is synonymous with aminosidine, which is marketed outside the United States. It is a poorly absorbed oligosaccharide aminoglycoside, related to neomycin, kanamycin, and other aminoglycosides of the catenulin group, which achieves high concentrations in the colon (Scaglia et al., 1994). The oral form has been available in the United States for many years for treatment of amebiasis, whereas the intravenous form is not available in the country because of its toxicity. Since 1990, when its use in the treatment of cryptosporidiosis in patients with AIDS was first reported (Gathe et al., 1990), there have been numerous reports of dramatic responses to this agent (Andreani et al., 1983; Armitage et al., 1992; Bissuel et al., 1994; Clezy et al., 1991; Danziger et al., 1993; Fichtenbaum et al., 1993; Wallace et al., 1993; Whiteside et al., 1984). Nonetheless, many patients continue to shed oocysts and others relapse while on therapy, suggesting that the agent is static and not cidal for Cryptosporidium, that a nonabsorbable agent may not be able to induce a complete and long-lasting response, or both. A placebo-controlled study that involved 10 patients randomized to paromomycin or placebo showed a decrease in both bowel movement frequency and oocyst shedding with paromomycin. Problems with the study include the small sample size and the persistence of a median of 109 × 106 oocysts in the stool of paromomycin-treated responders. Also worrisome is that four patients developed biliary tract disease and three required cholecystectomy during the follow-up period. A pilot study examining the safety and efficacy of paromomycin found a 50% reduction in oocyst excretion with 65% of cryptosporidiosis patients demonstrating complete or partial response (Blanshard et al., 1997). A study involving paromomycin given in combination with azithromycin reported a 95% reduction in oocyst excretion after 4 weeks and a 99% reduction after 12 weeks (Smith et al., 1998). A second study using the combination of paromomycin and azithromycin to treat an AIDS patient with respiratory cryptosporidiosis proved effective, with a complete response after 27 days of treatment (Meamar et al., 2006). Children have also been reported as responding well to paromomycin at 35mg/kg/day (Trad et al., 2003). In contrast, a prospective, randomized, double-blind, placebo-controlled study involving 35 AIDS patients with cryptosporidiosis demonstrated that there were no significant differences in efficacy between paromomycin and placebo (Hewitt et al., 2000). Despite conflicting data, controlled and uncontrolled, paromomycin is currently used widely as the first-line agent against cryptosporidial infection in patients with AIDS. Most patients enter treatment trials after paromomycin has failed. Whether delay in seeking other therapy increases their chances of failure, possibly because of spread to the biliary tree, is not known but worth considering. The role of paromomycin in treatment of cryptosporidiosis continues to be problematic.
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263
Nitazoxanide (Alinia®, Romark Laboratories)
This is a nitrothiazole benzamide compound first synthesized by Rossignol in 1976 with a wide spectrum of activity against protozoan, helminthic, and bacterial pathogens, including flagellates, coccidians, amoebae, nematodes, cestodes, and trematodes (Rossignol and Maisonneuve, 1984; Rossignol et al., 1984). In 2002, the U.S. Food and Drug Administration (FDA) approved nitazoxanide for the treatment of Cryptosporidium and Giardia intestinalis infections in children and, in 2004, approved it for the treatment of Giardia in adults (Fox and Saravolatz, 2005). In 2005, nitazoxanide was approved for treatment of Cryptosporidium infections in adults and teens (Freid, 2005). This drug is available in both oral and tablet formulations (Fox and Saravolatz, 2005), The recommended dosage for children between 12 and 47 months is 100 mg twice a day for 3 days, and 200 mg twice a day for 3 days for children 4 to 11 years old. Adults should take 500 mg twice a day for 3 days (Fox and Saravolatz, 2005; Freid, 2005). A randomized, double-blind, placebo-controlled study demonstrated that nitazoxanide had an 81% clinical cure rate and a 67% parasitologic cure rate 7 days after the initial treatment (Rossignol et al., 2001).
c.
Difluoromethylornithine (DFMO) (Eflornithine®, Ornidyl®, Sanofi-aventis)
This is an irreversible inhibitor of ornithine decarboxylase, which interferes with the biosynthesis of putrescine and other polyamines. Despite two reports of some success with the use of this agent in AIDS patients with cryptosporidiosis, serious toxicity (bone marrow suppression, gastrointestinal intolerance, and hearing impairment) has impeded any further development (Rolston et al., 1989; Soave et al., 1985).
d.
Atovaquone (Mepron®, Glaxo Wellcome)
This is an antimalarial hydroxy-naphthoquinone evaluated for activity against Pneumocystis carinii, Toxoplasma gondii, and Cryptosporidium in patients with AIDS. Results obtained in trials for cryptosporidiosis are not readily available but, reportedly, are disappointing (Gutteridge, 1991). There have also been anecdotal reports of improvement of cryptosporidial diarrhea in AIDS patients with other agents, including zidovudine (AZT) (Burroughs Wellcome) (Chandrasekar, 1987; Greenberg et al., 1989) and recombinant interleukin-2 (Kern et al., 1985). These effects are most likely due to augmentation of immune function, rather than a direct antiparasitic action.
C.
Highly Active Antiretroviral Therapy (HAART)
HAART was introduced in 1996 and has decreased the number of opportunistic viral, bacterial, and parasitic infections in HIV patients (Cacciò and Pozio, 2006; Palella et al., 1998; Pozio and Morales, 2005). It has become both a prophylactic and chemotherapeutic treatment for cryptosporidiosis in immunocompromised patients, with numerous cases reporting that the therapy can resolve cryptosporidiosis in most immunocompromised patients (Carr et al., 1998; Foudraine et al., 1998; Maggi et al., 2000; Miao et al., 1999; Moling et al., 2005; Schmidt et al., 2001). HAART increases CD4+ T-cell counts and inhibits viral replication using a combination of nucleoside and non-nucleoside reverse transcriptase inhibitors (NNRTIs) and HIV protease inhibitors (Table 9.2) (Cacciò and Pozio, 2006). In treating cryptosporidiosis with HAART, an NNRTI or protease inhibitor (PI) is normally used in conjunction with two or more nucleoside reverse transcriptase inhibitors (NRTIs) (Ives et al., 2001). Nucleoside analogs or NRTIs, target reverse transcriptase via competitive binding and inhibit DNA synthesis (Hoffmann and Mulcahy, 2006b). NNRTI also target reverse transcriptase, binding noncompetitively and blocking nearby active sites. This will slow DNA synthesis (Hoffmann and Mulcahy, 2006a). PIs inhibit HIV protease and proteolytic splicing, preventing the viral particles from becoming infective (Hoffmann and Mulcahy, 2006c). Cryptosporidiosis becomes chronic in immunocompromised patients with CD4+ T-cell counts lower than 180/µL (Carr et al., 1998) and life threatening in patients with CD4+ T-cell counts lower than 50/µL (Hoffmann, 2006). In these patients, improving the immune system to levels comparable to a healthy individual with HAART results in selflimiting diarrhea and resolution of cryptosporidiosis (Hoffmann, 2006). A study involving more than 3000 HIV-infected patients, over a 5-year period, undergoing treatment with antiretroviral combinations
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Cryptosporidium and Cryptosporidiosis, Second Edition TABLE 9.2 The Drug and Trade Names of NRTIs, NNRTIs, and PIs Used in Combination During HAART Therapy in Immunocompromised Patients Drug Name
Trade Name
Abacavir AZT-zidovudine ddC-zalcitabinea ddI-didanosine d4T-stavudine FTC-emtricitabine 3TC-lamivudine Tenofovir Nevirapine Efavirenz Delavirdine Amprenavir Atazanavir Fosamprenavir Indinavir Lopinavir Nelfinavir Ritonavir Saquinavir Tipranavir
Ziagen Retrovir HIVID Videx Zerit Emtriva; originally Coviracil Epivir Viread Viramune Sustiva or Stocrin Rescriptor Agenerase Reyataz Telzir or Lexiva Crixivan Kaletra Viarcept Norvir Invirase 500; originally Invirase or Fortovase Aptivus
Drug Class NRTI NRTI NRTI NRTI NRTI NRTI NRTI NRTI NNRTI NNRTI NNRTI PI PI PI PI PI PI PI PI PI
Note: Efficacies depend on status of the patient and the combination of drugs used. NRTI = nucleoside reverse transcriptase inhibitors; NNTRI = non–nucleoside reverse transcriptase inhibitors; PI = protease inhibitors. a
This drug was withdrawn from the market in June 2006.
Source: From Hoffmann, C., 2006. Cryptosporidiosis, in HIV Medicine 2006, 14th ed., Hoffmann, C., Rockstroh, J. and Kamps, B.S. (Eds). Flying Publisher, 825; Hoffmann, C. and Mulcahy, F., 2006a. Non-Nucleoside Reverse Transcriptase Inhibitors (NNRTIs), in HIV Medicine 2006, 14th ed., Hoffmann, C., Rockstroh, J. and Kamps, B.S. (Eds). Flying Publisher, 825; Hoffmann, C. and Mulcahy, F., 2006b. Nucleoside analogs (“nukes,” NRTIs), in HIV Medicine 2006, 14th ed., Hoffmann, C., Rockstroh, J. and Kamps, B.S. (Eds). Flying Publisher, 825; Hoffmann, C. and Mulcahy, F., 2006c. Protease inhibitors (PIs), in HIV Medicine 2006, 14th ed., Hoffmann, C., Rockstroh, J. and Kamps, B.S. (Eds). Flying Publisher, 825.
of NRTIs and PIs, showed a decreasing incidence of opportunistic infections and death. The incidence of cryptosporidiosis decreased from 0.8/100 patients to 0.1/100 patients (Moore and Chaisson, 1999).
D.
Supportive Therapy
Nonspecific antidiarrheal agents, including kaolin plus pectin (Kaopectate), loperamide (Imodium), diphenoxylate (Lomotil), bismuth subsalicylate (Pepto-Bismol), and opiates (tincture of opium, paregoric), are often helpful, but regimens must be individualized for each patient. The safety of nonspecific antidiarrheal therapy in patients with cryptosporidiosis is not known. A synthetic cyclic octapeptide analog of somatostatin, octreotide (Sandostatin, Sandoz Pharmaceuticals) has shown efficacy in patients with severe refractory secretory diarrhea caused by pancreatic cholera syndrome and carcinoid syndrome. A number of anecdotal reports suggest successful use of this agent in treatment of AIDS-related diarrhea (Clotet et al., 1989; Fanning et al., 1991; Moroni et al., 1993; Romeu et al., 1991). In a prospective, multicenter trial of escalating doses of subcutaneously administered octreotide in 51 AIDS patients, the response rate was 41%; most responders had neither cryptosporidial infection nor other identifiable pathogens (Cello et al., 1991). In a similar study of 34 AIDS patients in France, another somatostatin analog, vapreotide, was more likely to yield a response in patients with
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conditions other than cryptosporidiosis (Girard et al., 1992). Experience with intravenous somatostatin analogs is limited (Kuhls et al., 1994). Careful management of fluid and electrolyte balance is of paramount importance. Patients not able to maintain stable hydration and electrolyte balance may need intravenous replenishment and possibly nutritional support. The use of total parenteral nutrition is controversial because it requires an invasive procedure and is very expensive. Psychological support is frequently needed to help patients overcome the tremendous anxiety they experience due to fear of incontinence and, hence, the inability to move around freely.
E.
Future Directions
The search for efficacious therapies for human and animal cryptosporidiosis has been, for the most part, unrewarding (Georgiev, 1993; Ritchie and Becker, 1994; Soave, 1990). The FDA approval of nitazoxanide in humans has provided a means to treat this important parasite (Fox and Saravolatz, 2005; Freid, 2005). The introduction of HAART (Cacciò and Pozio, 2006; Palella et al., 1998; Pozio and Morales, 2005) has resulted in a decrease in chronic cryptosporidiosis in immunocompromised patients. These medical milestones provide a stimulus for new research leading to novel, and more efficient anticryptosporidial treatments. In patients in whom therapeutic intervention has failed, many of whom suffer from both AIDS and cryptosporidiosis, other factors such as profound immune dysfunction, multiple concomitant infections and therapies, and inadequate drug delivery to intracellular and perhaps biliary targets are likely responsible. Failure of therapeutic agents may also rest in the changing taxonomy of Cryptosporidium. Certain ultrastructural and biological analyses suggest that Cryptosporidium species have closer affinities to organisms other than those of the classical coccidia (Hijjawi et al., 2004; Tenter et al., 2002). This is further supported by the ability of C. parvum to successfully replicate extracellularly in hostcell-free media (Hijjawi et al., 2004). If Cryptosporidium species are not a coccidial organism, it may explain the failed attempts at treatment with anticoccidial therapeutic agents.
III. Treatment and Prophylaxis in Animals Numerous compounds have undergone efficacy evaluations against Cryptosporidium spp. in nonhuman hosts (Tables 9.3–9.5). Results were obtained from studies that focused on treatment of naturally acquired infections, or treatment or prophylaxis of experimentally induced infections or disease. Although both therapeutic and prophylactic drug regimens have been used, most efficacy evaluations of candidate agents consisted of the latter. Most studies evaluating potential anticryptosporidial agents were conducted in laboratory rodents, including inbred and outbred strains of mice, as well as laboratory rats and hamsters (see also Chapter 19). In many instances, laboratory species were rendered more susceptible to infection with Cryptosporidium by treatment with immunosuppressive agents such as hydrocortisone or dexamethasone, or consisted of genetically immunodeficient stock, such as nude or SCID mice. Many compounds show promise in laboratory rodents (Table 9.3), including the newly approved chemotherapeutic agent in humans nitazoxanide (Baishanbo et al., 2006; Blagburn et al., 1998; Gargala et al., 2005). The introduction of HAART has led to efficacy evaluations of antiretroviral therapy (Mele et al., 2003) and novel treatments involving monoclonal antibodies, and cytokines associated with the immune response (Gamra and Hosseiny, 2003; McDonald et al., 2004; Smith et al., 2001). In some instances, efficacies exceeded 90% compared to nontreated or diluent-treated controls. Treatment with other compounds resulted in modest but demonstrable reductions in parasite numbers compared to controls. Occasionally, infection relapsed when drugs were discontinued. Table 9.4 summarizes data derived from evaluation of potential anticryptosporidial agents in ruminants. Among drugs tested, paromomycin, lasalocid, halofuginone, and sulfaquinoxaline (Fayer, 1992; Fayer and Ellis, 1993b; Fayer et al., 1991; Fischer, 1983; Gobel, 1987a; Gobel, 1987b; Joachim et al., 2003; Lallemond et al., 2006; Mancassola et al., 1995; Moon et al., 1982b; Naciri et al., 1993; Naciri and
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TABLE 9.3 Efficacies of Potential Anticryptosporidial Drugs in Laboratory Rodents Animal
Age
C57BL6N mice Mature (immunosuppressed with dexamethasone) C57BL/6N 8 weeks (immunosuppressed with dexamethasone phosphate) C57/BL6 mice Infant
Drug
Efficacy
Reference
Dehydroepiandrosterone
10 mg/kg/day subcutaneously days 6–19
+
Rasmussen et al., 1995
Sodium selenite
12 µM Treatment with Se supplement on day 1 postinfection
+
Huang and Yang, 2002
Agmatine
32 mg/kg BID orally 2 days prior to infection until 5 days postinfection 32 or 64 mg/kg orally 5 h prior to infection and 5 h postinfection 3 mg/kg/day Administered in drinking water for 3 weeks 4 or 5 mg/mL in 250 µL volume given via gastric intubation beginning 35 or 39 days postinfection every 12 h for 21 days 0.25 g/kg/day 0.5 g/kg/day 1 g/kg/day 2 g/kg/day Orally for 10 consecutive days 50 mg/kg by I.P. injection every 12 h 10, 20 and 50 µM 10 µM (24 mg/kg) Treatment was given per os beginning at time of infection for 5 days or beginning 72 h postinfection for 3 days 100 μg given subcutaneously on the day of infection or 4 days postinfection 4% in diet for 14 days
+
Moore et al., 2001
+ +
Mead et al., 1995
+/–
Riggs et al., 2002
– – + +
Healey et al., 1995
–
Kuhls et al., 1994 Mele et al., 2003
C.B-17 SCID mice
Mature
Maduramicin Alborixin
C.B.17/Icr Tac-SCID mice
Mature
Monoclonal antibodies: 3E2(anti-CSL) 3H2(anti-GP25-200) 1E10(anti-P23)
C57BL/6N mice (treated with dexamethasone)
Mature
Paromomycin
BALB/c and SCID mice BALB/c mice
Mature
Aminoguanidine
3 days
PI cocktail (ritonavir, saquinavir, indinavir) Indinavir (alone)
Neonate
Mature
Rat anti-mouse IL-4 monoclonal antibody 11B11 L-arginine
Albino mice (immunosuppressed with hydrocortisone acetate)
2 weeks
Paromomycin
Athymic mice (intact and splenectomized)
Mature
BALB/cIL-4Rα-/-, IL-4-/-, IFN-γ-/mice Nude mice
Dosage and Method of Treatmenta
rIL-12
Lasalocid Sinefungin Dehydroepiandrosterone
100 mg/kg for 10 days administered orally 0.5 μg/mouse in 0.25 mL PBS for 3 days administered subcutaneously Treatments were given 1 day prior to infection and 3 days postinfection, alone and in combination 120 mg/kg/day for 3 days 20 mg/kg/day for 8 days 10 mg/kg/day for 8 days 120 mg/kg/day for 8 days
+ +
+
McDonald et al., 2004
+
Leitch and He, 1994a Gamra and Hosseiny, 2003
+
+ – – +/–
Leitch and He, 1994b
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TABLE 9.3 (CONTINUED) Efficacies of Potential Anticryptosporidial Drugs in Laboratory Rodents Animal BALB/c and SCID mice BALB/c mice
BALB/c mice
BALB/c mice
BALB/c-Ifgtml and C57BL/6- Ifgtml mice
Age
5–6 days and Clarithromycin 7 weeks 14-OH Clarithromycin Neonate Artemisinin β-Arteether β-Artemether Neonate Azithromycin Clarithromycin Erythromycin Paromomycin Neonate Sucrose Isomaltose N-acetyl-D-glucosamine Lactose Maltose Glucose Fructose 6–8 weeks rIL-12
Dosage and Method of Treatmenta
Efficacy
Reference
17.5–200 mg/kg
+/–
200 or 400 mg/kg administered subcutaneously or intrarectally 100 or 200 mg/kg orally prior to infection (0 h) and 24, 48, and 72 h after infection 5 g of sugar mixed with 10 mL distilled water given 24 h before infection or at the time of infection
– – – + +/– +/– + + – – – – – – –
Cama et al., 1994 Fayer and Ellis, 1994
0.5 μg intraperitoneally 1 day before infection and 0.25 μg/day for 7 days after rIL-4 infection 0.1 μg intraperitoneally beginning 1 day before rIL-4 plus rat IgG1 anti- infection, continuing for 7 IL-4 mAb days 0.4 μg rIL-4 plus 30 μg mAb intraperitoneally beginning 1 day before infection, continuing for 7 days
Neonate C57BL/6J, C57BL/6J- Ifgtml, BALB/cByJ and TLR9-deficient mice
CD-1 Swiss mice
Neonate
CD-1 Swiss mice
Neonate
Swiss ARC mice
Drug
Neonate
100 μg in 2.5 µL volume given orally 100 μg in 20 µL volume given intraperitoneally; treatments given 1 day before and 1 day after infection Diloxanide furoate 10 mg/kg β-Clycodextrin 34 mg/kg Treatments administered alone and in combination orally 1 h before infection and 1–5 days postinfection G1 (1-(5-bromofur-2-yl)- 20 mg/kg 2-bromo-nitro-ethene) βCD (beta-cyclodextrin) 833 mg/kg G1- βCD inclusion 20 mg/kg plus 833 mg/kg complex All treatments were administered orally 1 or 2 h before inoculation and every 12 h or 8 h days 1–5 postinfection Trifluralin 100 mg/kg given orally via Oryzalin gastric intubation for 4 days postinfection at 12 h intervals CpG oligodeoxynucleotides (ODNs)
Fayer and Ellis, 1993a
Harp, 1999
Smith et al., 2001
–
–
+
Barrier et al., 2006
+
CastroHermida et al., 2001
+
CastroHermida et al., 2004a
+ +
– +
Armson et al., 1999
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TABLE 9.3 (CONTINUED) Efficacies of Potential Anticryptosporidial Drugs in Laboratory Rodents Animal
Age
Drug
Swiss IRC mice
Neonate
Dicationic carbazoles (N = 18) Paromomycin Nitazoxanide
SCID mice
Neonate
Atovaquone
SCID mice
5 weeks
IgY (Chicken egg yolk antibody)
TCR-α-deficient mice 1 week
Outbred mice (immunosuppressed with prednisolone)
Mature
Outbred mice
3 days
Outbred mice
7 days
Inbred and outbred mice
1–3 days
Dosage and Method of Treatmenta 0.65–20 mg/kg 50 mg/kg 100 and 150 mg/kg Treatments were given orally beginning the day of infection to 5 days postinfection ≥ 100 mg/kg/day
25% Anti-C. parvum IgY powder (w/w) given in feed 20% anti-C. parvum IgY solution (v/v) given in water Treatment was administered for 17 days postinfection [1N,12N]Bis(Ethyl)-cis134 mg/kg BID for 2 or 7 days 6,7-Dehydrospermine 50 mg/kg BID for 10 days (polyamine analog: SL- 130 mg/kg BID for 7 days 11047) Putrescine Treatments were given orally in 100 µL volumes beginning 4 days prior to infection Lasalocid 64 mg/kg/day 128 mg/kg/day Azithromycin 400 mg/kg/day for 3 days Maduramicin 1.0, 2.5 mg/kg Alborixin 1.5, 2.5 mg/kg Enrofloxacin 1.0, 3.0 mg/kg Aromatic amidines 2.8, 11.3 mg/kg orally daily for 5 days Lasalocid 20–30 mg/kg daily for 5–7 days Clopidol Clopidol X 10 Methylbenzoquate Methylbenzoquate X 10 Clopidol + Methylbenzoquate Clopidol + Methylbenzoquate X 10 Robenidine HCl Robenidine HCl X 10 Decoquinate Decoquinate X 2 Furazolidone Furazolidone X 5 Amprolium Arprinocid Nicarbazin Dinitolmide Furaltedone Furaltedone X 10 Sulphaquinoxaline
Efficacy
Reference
+/–
Blagburn et al., 1998
–
Rohlman et al., 1993 Kobayashi et al., 2004
+
+/– –
Waters et al., 2000
+ + +
Kimata et al., 1991
+ + – +/–
Blagburn et al., 1991
+
Gobel and Bretschneider, 1985 Angus et al., 1984
0.25 mg/mouse/day 2.5 mg/mouse/day 0.25 mg/mouse/day 2.5 mg/mouse/day 0.1 mg/mouse/day
– – – – –
1.0 mg/mouse/day
–
0.03 mg/mouse/day 0.3 mg/mouse/day 2.0 mg/mouse/day 4.0 mg/mouse/day 0.4 mg/mouse/day 2.0 mg/mouse/day 0.25 mg/mouse/day 0.06 mg/mouse/day 0.125 mg/mouse/day 1.25 mg/mouse/day 0.2 mg/mouse/day 2.0 mg/mouse/day 2.8 mg/mouse/day
– – – – +/– + – +/– – – +/– + Toxic
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TABLE 9.3 (CONTINUED) Efficacies of Potential Anticryptosporidial Drugs in Laboratory Rodents Animal
Age
Drug Salinomycin Halofuginone
C57 mice
Neonate
Hamster
20–24 months
Hamster
5 days 12 days
Gerbil (immunosuppressed with dexamethasone)
?
Gerbil (immunosuppressed with dexamethasone)
1 month
Wister rat
Neonate
Rat
Neonate
Dosage and Method of Treatmenta
0.06 mg/mouse/day 0.003 mg/mouse/day Treatment began two days before infection and continued for about 7 days Ethopabate 28 mg/mouse/day Nicarbazin 0.1 mg/mouse/day Sulfaquinoxaline 30 mg/mouse/day Furaltadone 0.1 mg/mouse/day Enterolyte-N 0.02 mL/mouse/day Sulphamethazine 5 mg/mouse/day Trinamide 3 mg/mouse/day Amprolium 0.02, 8.0 mg/mouse/day Phenamidine 0.01 mL/mouse/day Zoaquin 0.5 mg/mouse/day Halofuginone 6 mg/mouse/day Salinomycin 0.02 mg/mouse/day Monensin 0.04 mg/mouse/day Emtryl 4 mg/mouse/day Arprinocid 8 mg/mouse/day Daily for 8 days after infection (Dose doubled for last 4 days) Dehydroepiandrosterone 120 Φg/kg/day subcutaneously for 7 days prior to infection Arprinocid 0.5 or 1.0 mg/animal on days 3, 7, 11, 14 or 6, 10, 14, 18 after infection RM-6427 200 or 400 mg/kg/day × 8 RM-6428 days (5,7-dihydroxyisoflavone 400 mg/kg/day × 12 days derivatives) Nitazoxanide 200 mg/kg/day × 12 days Paromomycin 100 mg/kg/day × 12 days Nitazoxanide 200 mg/kg/day orally × 12 Paromomycin days 100 mg/kg/day orally × 12 days Treatment began the day of infection Trifluralin 100 mg/kg given orally via Oryzalin gastric tubation for 4 days postinfection at 12 h intervals Actimel (Lactobacillus 200 µL orally 2 days prior to casei, L. bulgaricus, infection and 2 days Streptococcus postinfection to sacrifice thermophilus) VSL#3 (L. acidophilus, 50 µL (2.107 CFU) given orally 2 days postinfection to L. casei, L. plantarum, sacrifice L. bulgaricus, Bifidobacterium longum, B. breve, B. infantis, S. thermophilus)
Efficacy
Reference
Toxic
– – – – – – – – – – – – – – –
Tzipori et al., 1982
+
Rasmussen and Healy, 1992
+/–
+ +
+ +/– +
Kim, 1987
Gargala et al., 2005
Baishanbo et al., 2006
+ – +
Armson et al., 1999
–
Guitard et al., 2006
–
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TABLE 9.3 (CONTINUED) Efficacies of Potential Anticryptosporidial Drugs in Laboratory Rodents Animal Rat (immunosuppressed with dexamethasone)
Age
Drug
Mature
Halofuginone
Rat Mature (immunosuppressed with dexamethasone) Rat Mature (immunosuppressed with hydrocortisone)
Paromomycin
Rat (immunosuppressed with dexamethasone)
Mature
Rat Mature (immunosuppressed with hydrocortisone)
Rat Mature (immunosuppressed with hydrocortisone) Mature Rat (immunosuppressed with dexamethasone)
Paromomycin
Paromomycin Gentamicin Neomycin Kanamycin A Polymixin B Streptomycin Azithromycin Sulfadoxinepyrimethamine Quinicrine Trimethoprimsulfamethoxazole Bleomycin Elliptinium Daunorubicin Pentamidine α-Difluoromethylornithin diclazuril N-methyglucamine Vitamin A Sinefungin Lasalocid Metronidazole Sulfadimethoxine Sinefungin
Dehydroepiandrosterone
Dosage and Method of Treatmenta
Efficacy
37.5 Φg/kg/day 75 Φg/kg/day 150 Φg/kg/day 300 Φg/kg/day 600 Φg/kg/day 900 Φg/kg/day Drug was added to feed for 11 days 50 mg/kg/day 100 mg/kg/day 200 mg/kg/day 400 mg/kg/day 10 mg/kg/day 50 mg/kg/day 100 mg/kg/day Treated for 10 days 200, 400 mg/kg/day 200 mg/kg/day 200 mg/kg/day 200 mg/kg/day 200 mg/kg/day 200 mg/kg/day 400 mg/kg/day 3–60 mg/kg/day
+ – – – – – + –
? 6–250 mg/kg/day
– –
0.05 mg/kg/day 5 mg/kg/day 2 mg/kg/day 60 mg/kg/day 25, 100, 400 mg/kg/day
– – – – –
4 mg/kg/day 50 mg/kg/day 10,000, 25,000 IU/day 0.25, 2, 6, 10 mg/kg/day 2,10 mg/kg/day 25, 50 mg/kg/day 10, 100 mg/kg/day 0.01–10 mg/kg/day (curative) 0.01–10 mg/kg/day (preventive) 60, 120 mg/kg/day
Reference
– – + + + +
Rehg, 1995
– + + + – – +
Verdon et al., 1995
– – +/– +/– + + + + + +
Verdon et al., 1994
Rehg, 1994
Lemeteil et al., 1993
Brasseur et al., 1993 Rasmussen et al., 1992
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TABLE 9.3 (CONTINUED) Efficacies of Potential Anticryptosporidial Drugs in Laboratory Rodents Animal
Age
Rat Mature (immunosuppressed with hydrocortisone)
Drug
Sulfadimethoxine Bleomycin Lasalocid Metronidazole Mepacrine Elliptinium Daunorubicin Diclazuril Glucanthine α-difluoromethylornithine
Dosage and Method of Treatmenta Drugs given days 14–28 (curative) 60 mg/kg/day 0.05 mg/kg/day 10 mg/kg/day 25 mg/kg/day 10 mg/kg/day 5 mg/kg/day 2 mg/kg/day 4 mg/kg/day 50 mg/kg/day 400 mg/kg/day
Efficacy
Reference Brasseur et al., 1991
+ – + + + – – +/– +/– –
Drugs given days 4–10 (preventive) Spiramycin Pristinamycin Lincomycin Clindamycin Norfloxacin Paromomycin Mepacrine Mefloquine Pentamidine Metronidazole Lasalocid Sulfadimethoxine α-Difluoromethylornithine Sulfaquinoxaline + Vitamin K Sulfamethoxazole + Trimethoprim Sulfadoxine + Pyrimethamine Diclazuril Rat (immunosuppressed with dexamethasone)
Mature
Succinylsulfathiazole Sulfabenzamide Sulfacetamide Sulfachloropyridazine Sulfadiazine Sulfadimethoxine Sulfadoxine Sulfaguanidine Sulfamerazine
100 mg/kg/day 70 mg/kg/day 50 mg/kg/day 100 mg/kg/day 10 mg/kg/day 50 mg/kg/day 10 mg/kg/day 25 mg/kg/day 2 mg/kg/day 25 mg/kg/day 2 mg/kg/day 10 mg/kg/day 10 mg/kg/day 60 mg/kg/day 400 mg/kg/day
– +/– – – +/– – – +/– + +/– + + +/– + – –
160 mg/kg/day
–
250 mg/kg/day 50 mg/kg/day 4 mg/kg/day 0.2 mg/kg/day 3 mg/kg/day 360 mg/kg/day 240 mg/kg/day 120 mg/kg/day 360 mg/kg/day 250 mg/kg/day 120 mg/kg/day 160 mg/kg/day 120 mg/kg/day 200 mg/kg/day
– – – +/– – – – – – + – – +
Rehg, 1991a
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TABLE 9.3 (CONTINUED) Efficacies of Potential Anticryptosporidial Drugs in Laboratory Rodents Animal
Age
Drug
Rat (immunosuppressed with dexamethasone)
Mature
Sulfameter Sulfamethazine Sulfamethizole Sulfamethoxazole Sulfamethoxypyridazine Sulfanilamide Sulfanilic acid Sulfanitran Sulfapyridine Sulfasalazine Sulfathiazole Sulfisomidine Sulfisoxizole
Rat (immunosuppressed with dexamethasone)
Mature
Rat (immunosuppressed with dexamethasone)
Mature
Azithromycin Clarithromycin Erythromycin Oleandomycin Spiramycin Azithromycin Spiramycin
Rat (immunosuppressed with dexamethasone) Rat (immunosuppressed with dexamethasone)
Mature
Arprinocid
Mature
Lasalocid
Monensin Salinomycin
Rat (immunosuppressed with dexamethasone)
Mature
Sulfadimethoxine
Dosage and Method of Treatmenta 120 mg/kg/day 175 mg/kg/day 480 mg/kg/day 320 mg/kg/day 160 mg/kg 120 mg/kg 120 mg/kg/day 200 mg/kg/day 240 mg/kg/day 400 mg/kg/day 120 mg/kg/day 240 mg/kg/day 120 mg/kg/day Administered in food or water 50, 100, 200, and 400 mg/kg/day in feed for 11 days 200 mg/kg/day only 200 mg/kg/day Administered by gavage or in feed 1 h before infection through 11 days after infection 50 mg/kg 25 mg/kg 12.5 mg/kg In feed for 11 days 18 mg/kg/day 9.0 mg/kg/day 4.5 mg/kg/day 2.25 mg/kg/day 1.12 mg/kg/day 9.0 mg/kg/day 9.0 mg/kg/day Both therapeutic and prophylactic regimens used 120 mg/kg/day 80 mg/kg/day 40 mg/kg/day 20 mg/kg/day 10 mg/kg/day
Efficacy
Reference
+ + – – – – – – – – – – +
Rehg, 1991a
+/– +/– +/– +/– +/– + –
Rehg, 1991b
Rehg, 1991c
+ + –
Rehg and Hancock, 1990
+ + + + – – –
Rehg, 1993
+ + + + –
Rehg et al., 1988
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TABLE 9.3 (CONTINUED) Efficacies of Potential Anticryptosporidial Drugs in Laboratory Rodents Animal
Age
Rat Mature (immunosuppressed with dexamethasone
Drug Diethyldithiocarbamate
Amphotericin B Eflornithine Ivermectin Levamisole Thiabendazole Thalidomide
Diethyldithiocarbamate
Dosage and Method of Treatmenta Prophylactic daily for 11 days from 1 h before inoculation 900 mg/kg/day 600 mg/kg/day 300 mg/kg/day 150 mg/kg/day 75 mg/kg/day 37.5 mg/kg/day 20 mg/kg/day 3000 mg/kg/day 0.4 mg/kg/day 4 mg/kg/day 50 mg/kg/day 150 mg/kg/day Therapeutic daily from days 10–21 600 mg/kg/day Drugs given subcutaneously or in feed for all studies
Efficacy
Reference Rehg, 1996
+ + + + + – – – – – – – – +
Note: + = Demonstrable activity against C. parvum; – = No demonstrable activity against C. parvum; +/– = Partial activity, activity at one or more dosages for a compound, or activity for one or more compounds in a group. a
Both therapeutic (curative) and prophylactic (preventive) regimens used in some studies.
Yvore, 1989; Naciri et al., 1984; Nagy et al., 1984; Peeters et al., 1993; Villacorta et al., 1991; Viu et al., 2000), as well as (based on recent efficacy evaluations) α-cyclodextrin, β-cyclodextrin, and decoquinate, possessed demonstrable or partial activity against C. parvum infections in ruminant species (Castro-Hermida et al., 2004b; Castro-Hermida et al., 2001; Ferre et al., 2005; Lallemond et al., 2006; Mancassola et al., 1997; Moore et al., 2003). In 1999, halofuginone lactate (Halocur®, Intervet) was approved in countries other than the United States for treatment and prevention of cryptosporidiosis in calves (Anonymous, 1999; Thompson et al., 2005). The dosage for calves is dependent on weight. Calves weighing 35 to 45 kg received 8 mL SID for 7 days, whereas calves weighing 45 to 60 kg received 12mL SID for 7 days beginning 24 to 48 h after birth for a prophylactic effect (Anonymous, 1999). Therapeutic effects, decreasing the number of oocysts excreted in the feces, have been demonstrated in calves that are treated within the first 24 h after developing diarrhea (Anonymous, 1999; Thompson et al., 2005). In many cases, these data support results obtained in laboratory animal models. Table 9.5 summarizes drugs evaluated in miscellaneous animal species. Although little information is available on trials in animals other than rodents and ruminants, some success was achieved in treating cryptosporidiosis in felids and elapid snakes (Barr et al., 1994; Cranfield and Graczyk, 1994). Efficacy evaluations of paromomycin and enrofloxacin showed demonstrable activity against Cryptosporidium baileyi infections in chickens (Sréter et al., 2002), as did hyperimmune bovine colostrum (HBC) against Cryptosporidium serpentis in various snake species (Graczyk et al., 1998) and C. parvum in Savannah monitors (Graczyk et al., 2000). Spiramycin appeared somewhat effective in the treatment of naturally acquired Cryptosporidium infections in elapid snakes from a zoological park (Cranfield and Graczyk, 1994). In vitro efficacy evaluation of anticryptosporidial agents has confirmed the in vivo efficacies of agents such as maduramicin, paromomycin, and sinefungin, and has also identified additional agents such as colchicine and vinblastine (Table 9.6) (Arrowood et al., 1991, 1994; Favennec et al., 1994; Marshall and Flanigan, 1992; McDonald et al., 1990; Weist et al., 1993). Efficacies were demonstrated also for some small peptides, which function presumably by interacting with and lysing membranes of invasive
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TABLE 9.4 Efficacies of Potential Anticryptosporidial Drugs in Ruminants Animal
Age
Drug
Calf
Neonate
Paromomycin
Calf
Neonate
Halofuginone lactate
Calf Calf
Neonate Neonate
Sulfadimethoxine Halofuginone lactate
Calf
Neonate
Halofuginone lactate Sulfadimethoxine
Calf
4–5 days
Lasalocid-Na
Calf
1–10 weeks
Sulphadimidine
Calf
Up to 14 days
Amprolium Sulphadimidine Trimethoprim + Sulphadiazine Dimetridazole Ipronidazole Quinacrine Monensin Lasalocid
Calf
Lasalocid
Calf
2 days–3 months 8 days
Calf
1 day
Calf
Neonate
Bovine serum concentrate (BSC) L-glutamine Recombinant bovine interleukin-12 (rBoIL-12) Decoquinate
Dosage and Method of Treatmenta 100 mg/kg 50 mg/kg 25 mg/kg Twice daily in milk for 11 consecutive days 30 Φg/kg 60 Φg/kg 120 Φg/kg Days 2–8 after infection 5 gram bolus daily for 21 days 30 Φg/kg 60 Φg/kg 125 Φg/kg 250 Φg/kg 500 Φg/kg (in milk daily for 3–14 days) 2 mL/10 kg body weight (0.05 g halofuginone base/100 mL solution) 10 mL Given orally for 7 days 15 mg/kg (administered daily for 3 days) Therapeutic (200 mg/kg for three 3-day courses) Preventative (30 mg/kg for two 7-day courses) Preventative (40 mg/kg for 14 and 7 days at interval of 6 days 0.45 g/calf/day 5.0 g/calf/day 0.2 g/calf/day 1.0 g/calf/day 1.0 g/calf/day 1.0 g/calf/day 0.5 g/calf/day 0.2 g/calf/day 0.3 g/calf/day 0.03 g/calf/day Drugs administered twice daily throughout study 6–8 mg/kg Daily for 3–4 days 56.7 g BSC plus milk replacer or oral rehydration solution prior to and after inoculation 9 g/L given with or without BSC 4–10 mL (1540 ng/mL) given intraperitoneally or subcutaneously 1 day before infection up to 6 days postinfection 2 mg/kg bid given orally in milk replacer beginning the day of inoculation up to 28 days
Efficacy
Reference
+ +/– +/–
Fayer and Ellis, 1993b
– + +
Naciri et al., 1993; Peeters et al., 1993 Fayer, 1992 Villacorta et al., 1991
– – + + + + +/–
Joachim et al., 2003
+/– +
Gobel, 1987a
–
Fischer, 1983
– – – – – – – – – – + (toxic) –
Moon et al., 1982b
+
Gobel, 1987b
+
Hunt et al., 2002
– –
–
Pasquali et al., 2006
Moore et al., 2003
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TABLE 9.4 (CONTINUED) Efficacies of Potential Anticryptosporidial Drugs in Ruminants Animal
Age
Drug
Calf
7–10 days
Lamb
1 day
Lamb
3–10 days
Paromomycin
Lamb
1 day
β-Cyclodextrin
Goat
2–4 days
Paromomycin
Goat
?
Sulfaquinoxaline
Goat
Neonate
Sulfadimethoxine
Goat
1 day
Decoquinate
Goat
Neonate
α-Cyclodextrin
Goat
3 days
Decoquinate
Goat
Pregnant dam 4 years
Decoquinate
Camel
Halofuginone hydrobromide Decoquinate Halofuginone lactate
Amprolium
Dosage and Method of Treatmenta 100 μg/kg 2.5 mg/kg Treatments given orally days 1–7 0.5 mg/kg/day 3 or 5 days beginning 2 days after inoculation 100 mg/kg/day for 3 days 200 mg/kg/day for 2 days Treatment given orally beginning at the onset of diarrhea 500 mg/kg given orally 1–3 days of age and after the onset of diarrhea for 3 days 100 mg/kg twice daily (total daily dose) from day before infection through day 10 after infection 100 mg/kg plus vitamin K1 for 10 days 75 mg/kg daily for 5 days
Efficacy
Reference
+ –
Lallemond et al., 2006
+ (toxic?)
Naciri and Yvore, 1989
+
Viu et al., 2000
+
CastroHermida et al., 2001 Mancassola et al., 1995
+
+/– –
2.5 mg/kg /day for 21 days dissolved in milk or water and given orally via syringe beginning 3 days postinfection 500 mg/kg for 6 days orally beginning 4 h prior to infection
+
2.5 mg/kg/day BID for 21 days dissolved in milk and given orally via syringe 2.5 mg/kg/day for 21 days prior to kidding mixed with pelleted rations Pellets fed orally for 10 days
+/–
+
+/– –
Nagy et al., 1984 Naciri et al., 1984 Mancassola et al., 1997 CastroHermida et al., 2004b Ferre et al., 2005 Ferre et al., 2005 Fayer et al., 1991
Note: + = Demonstrable activity against Cryptosporidium spp.; – = No demonstrable activity against Cryptosporidium spp.; +/ = Partial activity, activity at one or more dosages for a compound, or activity for one or more compounds in a group; ? = data unavailable. a
Both therapeutic (curative) and prophylactic (preventive) regimens used in some studies.
sporozoites and merozoites. As in laboratory rodents, in vitro efficacy evaluations of antiretroviral therapeutic agents used in HAART show both demonstrable and partial activity against C. parvum (Hommer et al., 2003; Mele et al., 2003). Similar to the recommendation for humans, effective treatment of animals suffering from cryptosporidiosis may require oral or parenteral rehydration with fluids and electrolytes, in addition to antidiarrheals and attempted chemotherapy with putative anticryptosporidial drugs. Rehydration is particularly important in young animals and those immunocompromised or suffering from intercurrent disease. The latter should receive appropriate antibiotic therapy if bacterial copathogens are involved. Prevention of cryptosporidiosis in animals, again similar to recommendations for humans, is best achieved by eliminating contact with viable oocysts. This is difficult, particularly on farms and in zoos, given the resistance of oocysts to disinfectants. Prevention is based largely on knowledge of the biology, life cycle, and modes of transmission of Cryptosporidium spp. Infected animals should be quarantined in facilities that can be cleaned and disinfected. Contaminated fomites should be cleaned thoroughly or discarded. Animal care personnel should wear clothing that can be cleaned regularly. Clean food and
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TABLE 9.5 Efficacies of Potential Anticryptosporidial Drugs in Miscellaneous Animals Animal
Age
Drug
Pig
1 day
DL-α-difluoromethylornithine
Cat
6 months
Paromomycin
Chicken (C. baileyi)
7 days
Chicken (C. baileyi)
1 day
Lasalocid Maduramicin Monensin Narasin Salinomycin Semduramicin Halofuginone Salinomycin Lasalocid Monensin
Chicken (C. baileyi)
6 days
Garlic extract Diclazuril Toltrazuril
Chicken (C. baileyi)
6 days
Enrofloxacin Paromomycin
Quail
4–6 days
Turkey
7 weeks
Peacock
3–4 days
Black rat snake Yellow rat snake Corn snake Various snake species (8)a
Mature
Savanna monitors
Oxytetracycline Neomycin Furazolidone Amprolium Chlortetracycline Oxytetracycline HCl Spiramycin
Mature
Hyperimmune bovine colostrum
Mature
Hyperimmune bovine colostrum (HBC)
Dosage and Method of Treatment 0.38–1.25 g/kg Fed in milk daily for 10 days after pigs were shedding oocysts 165 mg/kg Daily for 5 days Drugs added to feed at 1× or 2× recommended level, alone or in combination with the antioxidant duokvin
3 mg/kg feed 60 mg/kg feed 75 mg/kg feed 110 mg/kg feed Daily 5 days before through 20 days after infection 7 mL/L 1 ppm 25 ppm 125 ppm 250 ppm Treatments were administered orally days 0–2 postinfection 50–100 mg/kg added to drinking water day 1 postinfection 333 and 666 mg/kg added to basal ration day 1 postinfection ? ? ? ? ? 200 ppm administered in water 80 mg/kg 3 doses (1 dose every second day) mixed in baby food HBC volume at 1% (0.8–31.7 mL) snake body weight administered by gastric intubation HBC volume at 1% lizard body weight administered by gastric intubation
Efficacy
Reference
–
Moon et al., 1982a
+
Barr et al., 1994 Varga et al., 1995
– – – – – – – – – –
Lindsay et al., 1987
+/–
Sréter et al., 1999
+
Sréter et al., 2002
+ – – – – – – +/–
+
+
Hoerr et al., 1986 Glisson et al., 1984 Mason and Hartley, 1980 Cranfield and Graczyk, 1994 Graczyk et al., 1998
Graczyk et al., 2000
Note: + = Demonstrable activity against C. parvum or C. baileyi; – = No demonstrable activity against C. parvum or C. baileyi; +/– = Partial activity, activity at one or more dosages of a compound, or activity for one or more compounds in a group; ? = data unavailable. a
Infected with C. serpentis.
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TABLE 9.6 Efficacies of Potential Anticryptosporidial Drugs or Compounds In Vitro Cell Line or Technique A-549 cell line (human lung carcinoma)
MDCK cell line (canine kidney) CaCo-2 cell line (human adenocarcinoma) CaCo-2 cell line (human adenocarcinoma)
Drug
Efficacy
Reference Giacometti et al., 1996
Azithromycin Clarithromycin Roxithromycin Spiramycin Atovaquone Pyrimethamine Minocycline Artemisin Dapsone Maduramicin
0.25, 0.5, 1, 2, 4, 8, 16, and 32 mg/L 0.25, 0.5, 1, 2, 4, 8, 16, and 32 mg/L 0.25, 0.5, 1, 2, 4, 8, 16, and 32 mg/L 0.25, 0.5, 1, 2, 4, 8, 16, and 32mg/L 0.02, 2, and 20 mg/L 0.005, 0.05, and 0.5 mg/L 0.2, 2, and 20 mg/L 0.02, 0.2 and 2 mg/L 0.1, 1, and 10 mg/L 0.5 Φg/mL Drug added to culture media for 24–168 h
+/– +/– +/– +/– – + + – – +
Sinefungin Diminazene Paromomycin
1 Φg/mL 10 Φg/mL 100 Φg/mL Drugs added to culture media for study period of 3 days Sporozoites incubated with anti-C. parvum IgY Treatment for 40 min then incubated with cells for 1 h 104 M 104 M Oocysts incubated with cells and media containing drugs 5 Φg/mL 50 Φg/mL 500 Φg/mL 5000 Φg/mL Excysted sporozoites incubated in media containing drug for 24 h
+ – +
Favennec et al., 1994
+
Kobayashi et al., 2004
+ +
Weist et al., 1993
– +/– +/– +
Marshall and Flanigan, 1992
Caco-2 cell line
IgY (Chicken egg yolk antibody)
HT29.74 cell line (human enterocyte)
Colchicine Vinblastine
HT 29.74 cell line (human enterocyte)
Paromomycin
Not applicable
Lytic peptides: Hecate-1 Shiva-10 Cecropin-b
BTFE cell culture system (bovine fallopian tube epithelial)
Dosage, Concentration, or Method of Treatment
Sodium selenite
10 Φg/mL 100 Φg/mL 100 Φg/mL Excysted sporozoites incubated with test article for 60 min Viability determined by vital dye staining 0.0 µM 1.5 µM 3.0 µM 6.0 µM 9.0 µM 12.0 µM Oocysts incubated with cells and media for 48 and 96 h
+ + –
+/–
Arrowood et al., 1994
Arrowood et al., 1991
Huang and Yang, 2002
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TABLE 9.6 (CONTINUED) Efficacies of Potential Anticryptosporidial Drugs or Compounds In Vitro Cell Line or Technique
Drug
Dosage, Concentration, or Method of Treatment
HCT-8 cell line (ileocaecal adenocarcinoma)
Paromomycin Antiretroviral protease inhibitors: Amprenavir Nelfinavir mesylate Ritonavir Saquinavir Indinavir sulphate
HCT-8 cell line (human colonic epithelial) BM cell line (bovine peripheral blood monocytes) HCT-8 cell line (human ileocecal adenocarcinoma)
Dihydroxyisoflavone derivatives Trihydroxydeoxy-benzoin derivatives Dihydroxydeoxybenzoine substituted derivatives PI cocktail (ritonavir, 10, 20, and 50 µM of drug incubated saquinavir, indinavir) with infected cells Indinavir (alone) 50 µM of drug incubated with cells 24 h before addition of sporulated oocysts 0.1, 0.5, 5, 10, 20, and 50 µM of drug incubated with cells and sporulated oocysts simultaneously 50 µM of drug added to infected cells every 24 h for 4 days Trifluralin 5 µL of varied concentrations less Oryzalin than 0.5% incubated with cells and oocysts for 72–120 h Amphotericin B, Amprolium 0.0064–20 Φg/mL Arprinocid Excysted sporozoites incubated in Chloroquine media containing drug for 24 h Cycloguanil Diclazuril Glycarbilamide Halofantrine Methyl benzoquate Monensin Nigericin Oxytetracycline Robenidine Proguanil Pyrimethamine Spiramycin Sulfaquinoxaline Venturicidin Zidovudine
HCT-8 cell line
L929 cell line (mouse fibroblast)
1000 mg/mL alone or in combination with paromomycin: 1000 μg/mL 500 μg/mL 200 μg/mL 33 μg/mL 25 μg/mL 20 μg/mL 10 μg/mL 2.5 μg/mL Excysted sporozoites incubated with cells in media with inhibitor for 2 or 48 h 0.2 mL of drug in media incubated with excysted sporozoites and cells for 48 h
Efficacy
Reference
+/–
Hommer et al., 2003
+/– (some toxicity)
Gargala et al., 2005
+ +
Mele et al., 2003
+/–
Armson et al., 1999
+/–
McDonald et al., 1990
Note: + = Demonstrable activity against C. parvum; – = No demonstrable activity against C. parvum; +/– = Partial activity, activity at one or more dosages of a compound, or activity for one or more compounds in a group.
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water must be provided. The latter is now known to be increasingly important in waterborne transmission to animals and between animals and humans. Rodents and wild mammals should be restricted from access to animal quarters. Neonatal mammals should receive adequate amounts of colostrum early in life. Mammals that do not suckle should be fed milk replacer and, perhaps, parenteral vitamins to increase their appetites. In general, methods of control of cryptosporidiosis in animals remain the same as those published previously (Angus, 1990; O’Donoghue, 1995). Additional evaluations of newer compounds that have proved effective in laboratory animals and in cell cultures should be conducted in farm, zoo, and companion animals, when possible, to identify those with therapeutic potential. Subsequently, a combination of hygienic practices, effective chemotherapy, and supportive measures should result in effective control of most outbreaks of animal cryptosporidiosis.
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10 Foodborne Transmission
Ynes R. Ortega and Vitaliano A. Cama
CONTENTS I. II. III.
Introduction .................................................................................................................................. 289 Confirmed Outbreaks ................................................................................................................... 291 Detection Methods ....................................................................................................................... 293 A. Fresh Fruits, Vegetables, or Prepared Foods .................................................................. 293 B. Detection of Oocysts in Beverages................................................................................. 295 C. Shellfish ........................................................................................................................... 295 IV. Possible Sources of Foodborne Contamination........................................................................... 297 A. Contaminated Raw Ingredients in the Fields ................................................................. 297 B. Food Processing .............................................................................................................. 297 C. Pest Infestations .............................................................................................................. 298 D. Food Handlers ................................................................................................................. 298 V. Inactivation ................................................................................................................................... 298 VI. Hazard Analysis and Critical Control Points (HACCP) and Other Management Regulations ............................................................................................................. 299 VII. Conclusions .................................................................................................................................. 299 References........................................................................................................................... 300
I.
Introduction
Foodborne outbreaks of cryptosporidiosis have been reported since the early 1980s, parallel to the surge of reports of Cryptosporidium in human populations. Cryptosporidium has been identified from several food commodities, mostly fruits, vegetables, and shellfish (Table 10.1). Because these products are frequently eaten raw, they are a probable vehicle of infectious oocysts. The outbreaks of cryptosporidiosis have been frequently associated with waterborne or person-toperson transmission. In recent years, the United States has seen an increased number of outbreaks associated with drinking or contact with recreational waters. The foodborne transmission of Cryptosporidium is well documented; however, these outbreaks are less frequently described. This underreporting may be explained by several factors, primarily by timing and confounding elements. There is usually a long elapse of time between exposure to the parasite and the onset of symptoms, which is later followed by the outbreak investigation. Because of this time lag, the potentially contaminated food products are no longer available for testing, and the investigation has to rely primarily on specimens of the affected people and the epidemiological investigation. A strong confounding factor is the close interaction between water and foods. Water is a likely source of food contamination during irrigation, throughout the processing, or in the preparation of foods. The isolation of Cryptosporidium from fresh vegetables and shellfish worldwide supports the role of water as a contaminating source of food products. However, other potential sources of contamination such as animal or human contamination cannot be completely ruled out. 289
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TABLE 10.1 Foodstuffs in Which Cryptosporidium Oocysts Have Been Isolated Food Type Vegetables
Shellfish clams
Country Costa Rica Peru Norway Costa Rica Norway
Lettuce, parsley, cilantro, blackberries Seed sprouts
Monge and Arias, 1996 Monge et al., 1996 Ortega et al., 1997 Robertson and Gjerde, 2001b Calvo et al., 2004 Robertson et al., 2002
Spain and Italy
Dosinia exoleta, Veneurupis pullastra, and V. rhomboideus, Venus verrocosa Dosinia exoleta, Veneurupis pullastra, and V. rhomboideus, Venus verrocosa Dosinia exoleta, Veneurupis pullastra, and V. rhomboideus, Venus verrocosa Chamelea gallina clams
Freire-Santos et al., 2000 Gomez-Couso et al., 2004 Gomez-Couso et al., 2003 Traversa et al., 2004 Fayer et al., 2003
Mytilus galloprovincialis
Freire-Santos, et al., 2000; Gomez-Bautista et al., 2000; GomezCouso et al., 2004 Lowery et al., 2001 Gomez-Couso et al., 2003 Graczyk et al., 2001 Graczyk et al., 1999 Chalmers et al., 1997 Izumi et al., 2004 Fayer et al., 1998, 1999, 2002 Fayer et al., 2003
Spain and EU countries
Oysters
Cockles
Meat/meat products Tripe Apple cider
References
Cilantro, lettuce Radish, carrot. tomato, cucumber Lettuce, mint, cilantro, parsley Lettuce, mung bean sprouts
Spain
Mussels
Items
Italy Eastern United States and Canada Spain
Northern Ireland Spain and EU countries
Mytilus edulis Mytilus galloprovincialis
Canada United States Ireland Japan United States
Zebra mussel (Dreissena ploymorpha) Bent mussel (Ischadium recurvum) Mytilus edulis Corbicula japonica Crassostrea virginica
Eastern United States and Canada Spain
Crassostrea virginica Ostrea edulis
Spain and EU countries
Ostrea edulis
Netherland Spain
Crassostrea gigas Cerastoderma edule
Spain and EU countries
Cerastoderma edule
Europe
Small ruminants, including sheep and goat Tripe Apple cider
United Kingdom Ohio
Freire-Santos et al., 2002; Gomez-Couso et al., 2004 Gomez-Couso et al., 2003 Schets et al., 2007 Gomez-Bautista et al., 2000 Gomez-Couso et al., 2003 Pepin et al., 1997 Anonymous, 1985 Blackburn et al., 2006
Note: Updated table based on Millar, B.C., Finn, M., Xiao, L., Lowery, C.J., Dooley, J.S., and Moore, J.E. 2002. Cryptosporidium in foodstuffs—and emerging etiological route of human foodborne illness. Trends Food Sci.Tech.13, 168–187.
Produce can become contaminated by contact with irrigation water. When such water is contaminated with human or animal feces, fresh produce becomes at risk of contamination with Cryptosporidium oocysts (Peng et al., 1997). It is well known that other enteric pathogens such as Vibrio can be acquired by ingestion of raw shellfish and can lead to mortality of vulnerable populations. Significant morbidity and mortality of
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marine mammals (sea otters) that feed on mollusks have been associated with Toxoplasma infection (Conrad et al., 2005). Viable and infectious Cryptosporidium oocysts have been identified in clams, oysters, and mussels, raising concerns of their potential role in human infection. These shellfish are capable of filtering large volumes of water and are able to retain viable oocysts in their gills and gastrointestinal tract (Fayer et al., 2003; Gomez-Bautista et al., 2000; Gomez-Couso et al., 2003, 2005; Graczyk et al., 1998a; Graczyk and Schwab, 2000). Ocean water can be contaminated with agricultural runoff from adjacent farms, where a large number of oocysts are being excreted by farm animals (Sischo et al., 2000). This chapter will focus on foodborne outbreaks, detection and inactivation methods of Cryptosporidium oocysts in food items, and potential sources of contamination. It will also cover management practices to prevent food contamination with Cryptosporidium spp.
II.
Confirmed Outbreaks
The national estimates of the burden, trends, and sources of specific foodborne diseases in the United States are determined by FoodNet. FoodNet was created in 1995 as a collaborative effort of the Centers for Disease Control and Prevention (CDC), the Food Safety and Inspection Service (FSIS) of the U.S. Department of Agriculture (USDA), the Center for Food Safety and Applied Nutrition (CFSAN) of the Food and Drug Administration (FDA), and the Early Intervention Program (EIP) under the Individuals with Disabilities Education Act. Currently, FoodNet is the principal foodborne disease monitoring component of the CDC’s emerging infections program. Baseline data for foodborne bacteria were collected in 1995, and in 1997 Cryptosporidium and Cyclospora were included in this surveillance program. When FoodNet was created, five states and a few counties in three other states were part of the program. Since then, more sites were included each year and currently ten states, representing 14% of the U.S. population, are part of this surveillance study. These include Connecticut, Georgia, Maryland, Minnesota, New Mexico, Oregon, Tennessee, and selected counties in California, Colorado, and New York. Although the cases reported in FoodNet might not be strictly associated with food as the primary transmission route, it provides an indication of the overall trend of the incidence of foodborne Cryptosporidium infections in the United States. In 1997, 468 cases of cryptosporidiosis were reported and two were fatal. In 2003, the incidence rate of cryptosporidiosis was 10.9 cases per million persons (n = 481) (Anonymous, 2004) with a mortality rate of 0.68 (n = 3). In 2004, 637 cases of cryptosporidiosis were reported, with an overall incidence of 13.2 cases per million persons, of which 5 died (Anonymous, 2005). In 2005, Cryptosporidium was reported in 1313 patients with an overall incidence of 2.95 cases per 100,000 persons (Anonymous, 2006). It is not clear, however, how many of the overall cases reported to the FoodNet each year were exclusively foodborne. The first well-documented foodborne outbreak of cryptosporidiosis was reported in central Maine in 1993. Of the students and staff attending a school agricultural fair, 54% (154 individuals) acquired cryptosporidiosis. The epidemiological investigation indicated a strong association between infection and the consumption of hand-pressed apple cider, with a relative risk value of 26 (95% CI, 12–59). Subsequently, oocysts were detected in the apple cider, the cider press, and a calf from the farm that provided the apples (Millard et al., 1994), confirming the foodborne nature of this outbreak. In 1995, the Minnesota Health Department reported that approximately 50 persons who attended a social event acquired cryptosporidiosis. The investigation showed that consumption of chicken salad was associated with an increased risk of illness. Further investigations revealed that the chicken salad was prepared by the hostess, who also operated a licensed day-care facility. She reported that neither she nor any child attending the day-care facility had diarrheal illness; however, she also indicated that she had changed a child’s diapers and washed her hands on the day she prepared the chicken salad. Although there was no laboratory confirmation of the source of Cryptosporidium, this investigation suggested that asymptomatic children with cryptosporidiosis could be the source. Although the process of changing a diaper might have seemed routine and of little consequence, the closely timed activities at the day-care facility followed by preparation of food with contaminated hands or utensils were the only potential explanations for this outbreak (Anonymous, 1996).
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In 1996, during the second week of October, a total of 20 confirmed and 11 suspected cases of cryptosporidial diarrhea were identified in 19 households in New York State. People reportedly had diarrhea (100%), abdominal cramping (55%), vomiting (39%), fever (36%), and bloody diarrhea (10%). A case-control investigation revealed a history of drinking cider from a specific mill among case households, compared with only 1 of the 18 control households. The matched odds ratios (OR) for drinking cider pressed during September 28–29 were too high to be defined (p < 0.01). Reportedly, cider was made only from picked apples, and dairy livestock were not maintained by the orchard that supplied the apples, although a dairy farm was located across the road. Despite the epidemiological evidence, Cryptosporidium could not be detected in the remaining cider samples from the time of the outbreak, swabs of equipment surfaces from the mill, or water obtained on October 21 from the well in the cider mill (Anonymous, 1997). In October 2003, a northeast Ohio health department identified 12 local residents with laboratoryconfirmed cryptosporidiosis; 11 had drunk a locally produced, ozonated apple cider within 2 weeks preceding the illness. The remaining cider was embargoed, and the outbreak was investigated using epidemiologic and molecular techniques. Two separate case-control studies were conducted. In the first study, 12 of 19 case patients, but 0 of 38 controls, had drunk apple cider, and the matched OR for this association was incalculably high. However, drinking cider was the only variable associated with illness in a conditional logistic regression model (estimated OR, 14.0; 95% CI, 1.8–167). In the second study, 33 (10%) of the 329 persons who drank cider became ill, whereas only 2 (3%) of 73 who did not (adjusted relative risk, 4.7; 95% CI, 1.2–18.1). Samples from 14 laboratory-confirmed cases, and the remaining contents of cider drank by cases were tested using molecular tools for the species identification and subtyping of Cryptosporidium. Twelve samples (85.7%) were polymerase chain reaction (PCR) positive. Cryptosporidium parvum was identified in 11 of these, as well as in the sample of cider, whereas the remaining sample showed Cryptosporidium cervine genotype. The sample of cider had C. parvum subtype IIaA17G2R1, whereas the human isolates yielded two closely related subtypes: IIaA17G2R1 and IIaA15G2R1. The epidemiological investigation and the detection of C. parvum subtype IIaA17G2R1 in the sample of cider from a laboratory-confirmed case patient provided strong evidence that linked the consumption of cider with infected people. This outbreak highlighted the importance of the use of molecular tools in conjunction of epidemiological investigations to study Cryptosporidium outbreaks (Blackburn, 2006). Additional studies also suggested that other types of foods could be involved in the transmission of Cryptosporidium. In 1985, a retrospective study of 22 Canadian travelers with cryptosporidiosis revealed that the likely source of contamination was the consumption of nonpasteurized cow milk (Elsser et al., 1986). In Australia, two persons acquired cryptosporidiosis by drinking nonpasteurized goat milk (Table 10.2), and in 2002 eight additional people acquired cryptosporidiosis after drinking nonpasteurized cow milk (Harper, 2002). In September and October 1998, an outbreak of gastroenteritis affected 88 students and 4 cafeteria workers on a university campus in Washington, DC. The infectious agent was thought to be a virus until Cryptosporidium was identified in the feces of sick cafeteria employees. Although an additional 60 students reported gastrointestinal illness, data for these students were incomplete. The epidemiological investigation implicated two meals from one cafeteria as the likely sources of the outbreak. A food handler had the earliest onset of illness—characterized by copious watery diarrhea—but continued to work, preparing fruits and vegetables. Four days before the onset of illness, this employee had cared for an 18-month-old niece who had diarrhea and vomiting. This investigation was the first demonstration of the role of handlers in the contamination of food products and highlighted the importance of identifying index cases during a foodborne outbreak. The availability of molecular tools to discriminate species and genotypes greatly enhances the ability to perform trace-back studies and determine the source and route of transmission. In the case of the 1998 cafeteria outbreak, more than 100 persons were infected by an ill food handler, clearly demonstrating that Cryptosporidium is highly infectious and demonstrating the advantages of using molecular tools during investigations. This outbreak also shows that despite the existence of policies that food handlers should not work while ill, these are not strictly enforced (Quiroz et al., 2000).
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TABLE 10.2 Foodborne Outbreaks of Cryptosporidiosis Implicated or Suspected Food Raw goat milk
Number of Cases 2
Australia
Frozen tripe Sausage
1 19
United Kingdom Wales, U.K.
Milk
22
Mexico
154
Maine
Apple cider— unpasteurized Chicken salad Apple cider— unpasteurized Bovine milk—pasteurized Green onions Fruit/vegetables Unknown Unknown Unpasteurized milk
Ozonated apple cider
15 31 50 54 148 24 88 8
23
Suspected Mode of Transmission
Place
Minnesota Connecticut and New York United Kingdom Washington, D.C. Washington, D.C. Wisconsin Washington, D.C. Sunshine Coast, Queensland (Australia) Ohio
Consumption of unpasteurized goat’s milk Epidemiological association Consumption of milk by Canadian travelers Consumption of cider made of dropped apples Ill food handler Well water used for washing the apples Faulty pasteurizer
References Anonymous, 1984 Anonymous, 1985 Casemore et al., 1986 Elsser et al., 1986 Millard et al., 1994 Anonymous, 1996 Anonymous, 1997
Drank unpasteurized cow’s milk
Gelletlie et al., 1997 Anonymous, 1998 Quiroz et al., 2000 Millar et al., 2002 Millar et al., 2002 Harper et al, 2002
Once- and twice-ozonated apple cider
Blackburn et al., 2006
Unwashed green onions Ill food handler Company and private home
Note: Table adapted from Millar, B.C., Finn, M., Xiao, L., Lowery, C.J., Dooley, J.S., and Moore, J.E. 2002. Cryptosporidium in foodstuffs—and emerging etiological route of human foodborne illness. Trends Food Sci.Tech.13, 168–187.
III. Detection Methods Methods to detect Cryptosporidium oocysts in food products have been tested by many scientists. Generally, the following steps are used: elution of oocysts from the food, recovery and concentration of the oocysts, and detection. These steps can vary according to the food in question. For example, for detection of oocysts in liquid foods elution is not needed, and the existing protocols for detecting parasites in water have been applied.
A.
Fresh Fruits, Vegetables, or Prepared Foods
Methods have been developed to remove oocysts from leafy greens using elution buffers. One protocol includes washing fresh produce with distilled water by manual agitation for 10 min. The resulting washes are concentrated by centrifugation at 1000 × g for 15 min. The oocysts were either stained by modified acid-fast stain or fluorescent antibody (FA) and detected by microscopy. Using this method, oocysts were found in 14.6% of fresh vegetables purchased in markets serving endemic areas (Ortega et al., 1997). Oocysts have also been recovered from experimentally contaminated lettuce using a washing solution composed of sterile distilled water, 0.1% Tween 80, 0.05% sodium dodecyl sulfate (SDS), and antifoam A, at a ratio of 1:25, weight/volume of lettuce to the solution. The lettuce-wash buffer suspension was agitated in an orbital shaker for 5 cycles of 10 min each at 100 rpm. Oocysts were concentrated by filtration using Envirocheck capsules followed by adherence to immunomagnetic separation (IMS) beads as described for U.S. EPA Method 1623 (U.S. Environmental Protection Agency, 2001), then stained by modified Ziehl–Neelsen stain, and examined by microscopy. Recovery rates ranged from 0 to 6.5%.
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Detection limits using nested PCR for the cryptosporidium oocyst wall protein (COWP) gene were about 15 oocysts for concentrates before IMS and about 260 oocysts from glass slides (Ripabelli et al., 2004). Pulsification in glycine buffer also has been used to elute a small number of oocysts from lettuce. Oocysts were concentrated by centrifugation at 4000 × g for 30 min with recovery efficiencies up to 70% (Cook et al., 2006b). Oocysts were experimentally recovered from iceberg lettuce, bean sprouts, and strawberries using a wash solution composed of 150 mL of water and 50 mL of elution buffer described in U.S. EPA Method 1623. The buffer contained 1% (volume/volume) Laureth 12, 0.01-M Tris-HCl pH 7.4, 1-mM ethylenediamine tetra-acetic acid, disodium salt dihydrate (EDTA, 2 Na), pH 8.0, and 0.00015% Antifoam A (U.S. Environmental Protection Agency, 2001); 50 to 109 g of the produce, soaked in 200 mL of the solution, were washed twice by rotation in a drum for 1 to 5 min, followed by sonication for 3 min. The wash solutions were concentrated by centrifugation at 1000 × g for 10 min, and the pellets resuspended in 20 mL of distilled water. Half of the suspension was used to detect ova and oocysts. The remaining 10 mL were concentrated by centrifugation to 0.5 to 1.5 mL (Robertson et al., 2000), followed by IMS and FA detection of oocysts described in U.S. EPA Method 1623 (U.S. Environmental Protection Agency, 2001). The mean recovery efficiencies of this method ranged from 18 to 38% for iceberg lettuce, 1 to 5% for bean sprouts, and 12 to 35% for strawberries (Robertson and Gjerde, 2000). Cryptosporidium contamination was found in 5 of 125 samples of lettuce and 14 of 149 samples of mung bean sprouts (Robertson and Gjerde, 2001b). Recovery efficiencies with this method were higher when using 30 g of lettuce vs. 100 g, or 1-week-old mung bean sprouts vs. 10day-old noncontinuously refrigerated sprouts. Further evaluation of this method was done by direct spiking of Cryptosporidium into the wash solution and comparing the use of the IMS kit produced by Dynal (anti-Cryptosporidium kit) versus the kit produced by ImmuCell (CryptoScan kit). These modifications did not significantly change the recovery efficiencies of the method (Robertson and Gjerde, 2001a). Phosphate-buffered saline (PBS) or an elution solution of 1-M glycine and four different removal protocols were also used for oocyst recoveries. Either 30 g of lettuce or 60 g of raspberries were spiked with 100 Cryptosporidium oocysts. The samples were placed either in a 250-mL beaker with the glycine buffer and agitated with an orbital shaker at 80 rpm for 1 min or washed in a 500-mL plastic centrifuge pot with a spiral roller for 1 min. The eluates from these methods were concentrated by centrifugation at 2500 × g for 10 min, using 15ºC for lettuce and 4ºC for raspberries to minimize damage to the produce. Oocysts were removed from the pellets using IMS and examined with fluorescence microscopy along with 4′,6-diamidino-2-phenylindole (DAPI) stain for nuclei visualization (Smith et al., 2002). Recovery rates for lettuce were 24.8% from orbitally shaken samples and 36.4% from spirally rolled samples. The rates for raspberries were 40.5 and 45.4%, respectively. Similar lettuce samples were placed in Stomacher® bags with 200 mL of glycine-based elution buffer for 1 min to test the efficacy of Stomacher and the rapid pulse-beating mechanisms. The recovery rates from the spiked lettuce were 46.4 to 51.9% for Stomacher and 39.5 to 75% for the pulse-beating mechanism (Cook et al., 2006a). Other elution solutions were tested for compatibility with the CryptoScan IMS kit (ImmuCell Corporation). Phosphate-buffered saline with 0.1-M tricine at pH 5.4 or the elution buffer from U.S. EPA Method 1623 had low recovery rates, whereas solutions of 1-M sodium bicarbonate at pH 6.0, 0.1-M HEPES at pH 5.5, and 1% lauryl sulfate were not compatible with the CryptoScan IMS kit. In contrast, 1-M glycine at pH 5.5–5.6 was the most efficient elution buffer to be used with this kit (Cook et al., 2006a). Additional methods for leafy greens are also described in the literature. Samples of 200 g of cabbage or lettuce were spiked with 200 oocysts, placed in 1.5 L of an elution solution (1% SDS and 0.1% Tween 80), and processed in a sonic cleaning bath for 10 min. Oocysts were concentrated by centrifugation for 15 min at 1500 × g, and the resulting pellets were examined by microscopy. Those samples that produced large pellets were further purified over Sheather’s sugar solution and centrifuged at 1500 × g for 15 min. This method had a recovery efficiency of 1% for both cabbage and lettuce (Bier, 1991). An alternative procedure for indirect detection of Cryptosporidium has also been described. Antibodies specific to the capsid protein of a double-stranded RNA Cryptosporidium parvum virus (CPV) isolated from Cryptosporidium were used to detect oocysts from cilantro and green onions (Kniel and Jenkins, 2005). It is estimated that approximately 500 C. parvum symbiont viral particles or CPV are present per sporozoite. Cilantro and green onions were spot-inoculated with 106, 105, or 10 oocysts. The spiked
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samples were allowed to air-dry prior to elution. Oocysts were eluted from the cilantro and green onion samples by a sterile phosphate-buffered saline and 45 min of shaking at 100 rpm on an elliptical shaker (Kniel and Jenkins, 2005). The wash was concentrated by centrifugation at 4000 × g for 10 min, and the resulting pellet was resuspended in 1 mL of PBS; 100 µL of the suspension was tested by immunoblot, reacted with rabbit anti-CPV antiserum, followed by incubation with goat antirabbit biotinilated IgG, and visualized using avidin-alkaline phosphatase. Reportedly, this method detected as few as 10 oocysts.
B.
Detection of Oocysts in Beverages
Methods for detecting Cryptosporidium in beverages are similar to those for the recovery and detection of parasites in several types of water. Recovery methods for Cryptosporidium from beverages are usually based on the U.S. EPA Method 1623 (U.S. Environmental Protection Agency, 2001) which uses filtration followed by elution, concentration by IMS, and detection by FA. Oocysts detected in 50 mL of apple cider can be also concentrated by Sheather’s sucrose gradient centrifugation (Deng and Cliver, 2000). Cryptosporidium oocysts detected by this method can be further characterized using molecular methods, such as PCR-RFLP based on the small subunit of the rRNA gene (Jiang et al., 2005). For molecular testing, DNA can be obtained from the oocysts by subjecting the sample to six consecutive cycles of freezing in liquid nitrogen for 5 min and thawing at 65ºC for 5 min, followed by DNA purification using the QiAamp DNA Mini Kit (Qiagen). This method was used to test production water and washed apples used in the manufacturing of apple cider, as well as fresh and finished cider from five Canadian producers. Cryptosporidium was detected in 2 of 111 water samples, 1 of 114 washed apple samples, 5 of 113 fresh cider samples, and 2 of 113 samples of finished cider (Garcia et al., 2006). PCR amplification based on the CpR1 was employed and DNAj-like protein genes were evaluated for detection of Cryptosporidium in raw milk. Cryptosporidium DNA from 17 different isolates was used to spike milk samples. These samples were treated with 1 mL of reconstituted Bacto-trypsin and 5 mL of Triton X-100, incubated for 30 min at 50ºC, and centrifuged at 5000 × g for 10 min. The resulting pellet was resuspended to 100 µL, and DNA was extracted with a lysing buffer containing 120mM NaCl, 10-mM EDTA, 25-mM Tris(pH 7.5), 1% sarcosyl, and proteinase K to a final concentration of 5 mg/mL. Oocysts were ruptured by 5 to 10 freeze–thaw cycles, followed by a 1-h digestion at 55ºC with proteinase K. The extracted DNA was amplified using primers for a 358-base fragment of the CpR1 gene (GenBank accession number M95743) or a 452-base fragment of the DNAj-like protein that is conserved among several Cryptosporidium species, including C. hominis, C. parvum, and C. meleagridis. The resulting PCR products were verified by DNA probe hybridization specific to Cryptosporidium. The CpR1-based PCR method detected DNA from 8 of 9 human isolates and 6 of 6 bovine isolates but did not amplify C. muris. In contrast, the DNAj-based PCR amplified DNA from 5 of 9 human isolates, 3 of 6 bovine isolates, and from Eimeria acervulina. However, the detection limits for Cryptosporidium oocysts in raw milk using either of these PCR protocols was not determined due to inconsistent PCR amplification (Laberge et al., 1996).
C.
Shellfish
Oysters, clams, mussels, and other shellfish are filter feeders that remove particulates from the water they live in. Many shellfish live in waters that receive runoff from rural, suburban, and urban surfaces as well as industrial wastes, sewage discharges, and agricultural contamination. Shellfish, such as Corbicula fluminea can filter 2.5 L of water per hour, including suspended particles as large as 10 µm in diameter. Oocysts of Cryptosporidium have been identified in gill washings, hemolymph, and the digestive tract of shellfish (Fayer et al., 1998, 1999; Graczyk et al., 1998a, 1999). To detect parasites in gills or the gastrointestinal tract, tissues are excised after opening or shucking the shellfish. To obtain hemolymph, a hole can be drilled in the shell with an ordinary electric shop drill, and an 18-gauge needle fitted to a 5-mL syringe inserted through the hole into an adductor muscle from which hemolymph can be aspirated. Under controlled contamination conditions, Cryptosporidium parasites were observed in hemolymph, gills, and gastrointestinal tissues of clams as early as 1 day post exposure, with peak detection on day
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3 postexposure, and oocysts were still detectable at day 14 postexposure (Graczyk et al., 1998b). Tissues have also been processed by homogenization using an Ultra-Turrax T8 shearing-homogenizer or an Osterizer pulse-matic, with lipid extraction using a solution of diethyl ether and PBS (0.04 M, pH 7.2), followed by centrifugation at 1000–1250 × g for 5 min (Freire-Santos et al., 2000; Gomez-Couso et al., 2003) or by vortexing for 15 s in a Vortex-Genie (Scientific Industries, Bohemia, NY) set at the highest setting (Fayer et al., 2002, 2003). Other investigators recovered oocysts from the digestive tracts and gills using a wrist action shaker in 0.01-M PBS (pH 7.4). The gill tissues were then sieved through a 70-µm mesh nylon strainer and centrifuged at 1170 × g for 10 min. In other instances, digestive tract tissues were homogenized by squeezing and rubbing the tissues in 2.5-mL PBS in a small plastic bag, followed by sieving as described for gill tissues (Schets et al., 2007). Improvements on oocyst recovery rates include the use of IMS, even though this procedure requires the introduction of additional washing steps. Shellfish homogenates were centrifuged at 1000 × g for 15 min and the pellet was resuspended in reagent water and processed by IMS using Dynabeads (Invitrogen, Carlsbad, CA) as per the manufacturer’s instructions. The oocysts were dissociated from the immunomagnetic beads using 0.1N HCl and the sample was neutralized with 1N NaOH, followed by FA microscopy using fluorescein isothiocyanate (FITC)-labeled monoclonal antibodies (TCS Microbiology, North Buckinghamshire, United Kingdom) (MacRae et al., 2005). The recovery of oocysts from different tissues was compared using the IMS and FA methods. Whole shellfish homogenates had higher recovery rates than gill homogenates, gill washings, and hemolymph, with rates of 12 to 34% for mussels, 48 to 69.5% for oysters, and 30 to 65% for scallops (MacRae et al., 2005). The immunofluorescence assay provides a morphological identification of Cryptosporidium species, but this assay will not allow species determination. A fluorescent in situ hybridization assay (FISH) used C. parvum-specific oligonucleotide probes and labeled with a Hex red single fluorochrome tag on the 5′ end (Graczyk et al., 2004) that targeted the 18sRNA gene (Graczyk et al., 2003). FISH has been used in combination with FA to visualize both the oocyst wall and the nuclear contents. The samples (homogenates or washes) were fixed onto glass slides, treated with 70% acetic acid, and 5% dodecyltrimethyl ammonium bromide for 30 min at 80˚C to increase permeability (Graczyk et al., 2003). Samples were then used for FISH staining followed by FA. Using the same microscopy excitation filters as for FA, oocyst walls were visualized as apple green, whereas the nuclei were stained a reddish orange (Graczyk et al., 2003, 2004). For molecular analysis, Cryptosporidium DNA has been extracted using various methodologies. Hemolymph or gill washings were subjected to five freeze–thaw cycles, followed by phenol-chloroform extraction. The extracted DNA was evaluated using small subunit (SSU) rRNA-based PCR restriction fragment length polymorphism (RFLP). DNA also has been extracted using other DNA extraction kits; among them are the QIAamp DNA minikit (Qiagen Valencia, CA) or FASTDNA Spin KIT for Soil (MP Biomedicals, Irvine, CA) (Jiang et al., 2005). The SSU rRNA-based PCR-RFLP molecular tool has been used extensively for Cryptosporidium species or genotypes identification (Xiao et al., 1998). Analysis of 65 pooled oyster samples detected Cryptosporidium in 26 samples; 24 were C. parvum, 1 was C. baileyi, and the other was C. serpentis (Xiao et al., 1998). These results allow determination of the potential source of contamination. In this case, the most likely source of contamination was animals, as C. parvum is a parasite more frequently detected in young cattle (Feng et al., 2007; Santin et al., 2004). Briefly, the SSU rRNA based PCRRFLP method used 1 µL of extracted DNA for the nested PCR amplification. Each reaction contained 100 nM (primary PCR) or 200 nM (secondary PCR) primers, 0.2-mM deoxynucleoside triphosphate (Perkin-Elmer, Foster City, CA.), 3-mM MgCl2, 400 ng/µL of nonacetylated bovine serum albumin (Sigma, St. Louis, MO), 1X PCR buffer (Applied Biosystems, Foster City, CA), and 2.5U Taq DNA polymerase (Promega, Madison, WI) in a reaction volume of 100 µL. Negative controls consisting of PCR mixture without DNA template were included in each reaction. Primary and secondary PCR amplifications had a denaturation cycle of 94ºC for 2.5 min, followed by 35 cycles of 94ºC for 30 s, 55ºC for 60 s, and 72ºC for 3 min, with an additional 10 min extension at 72ºC. The amplification of Cryptosporidium DNA resulted in products of ~830 bp that were visualized in 2% agarose gels stained with ethidium bromide. For species and genotype identification, secondary PCR products positive for Cryptosporidium were digested overnight at 37ºC with restriction enzymes Ssp I and Vsp I. The digested
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products were electrophoresed in 2% agarose gels and the resulting banding patterns allow the identification of Cryptosporidium genotypes (Jiang et al., 2005; Xiao et al., 1998). See Chapter 5 for the protocol for the SSU rRNA-based PCR-RFLP genotyping tool.
IV.
Possible Sources of Foodborne Contamination
Cryptosporidium oocysts have been identified in a variety of food matrices, including vegetables and shellfish (Table 10.2). Fresh produce might be contaminated on the farm, shellfish might be contaminated when filtering oocysts from the water, or contamination might occur during food preparation by food handlers.
A.
Contaminated Raw Ingredients in the Fields
C. parvum, the most widespread zoonotic species, is a ubiquitous parasite frequently excreted by preweaned calves (Fayer et al., 2000; Santin et al., 2004). Other domesticated animals can be infected with other species of Cryptosporidium and, occasionally, C. parvum, which are excreted in animal feces and can contaminate agricultural fields. Proximity of dairy farms to agricultural fields may allow introduction of oocysts via contamination of irrigation water. The widespread practice of disposal of animal manure on land and its dispersal by runoff to rivers, lakes, and oceans is a concern as surface water can be contaminated with this runoff or by accidental contamination from human sewage (Peng et al., 1997). Additionally, a study of oocyst transport and subsurface irrigation methods demonstrated the presence of oocysts in soil at different depths (Armon et al., 2002). Fomites, including insects such as flies and dung beetles (Mathison and Ditrich, 1999; Szostakowska et al., 2004), and free-living nematodes (Huamanchay et al., 2004) may carry and disperse Cryptosporidium onto the vegetable surfaces. Wind or transport on shoes, clothing, vehicles, or tools can also contribute to contamination. Wildlife and free-living birds, such as gulls, might also play a role in parasite transmission (Smith et al., 1993). However, Cryptosporidium species infecting wildlife appear quite host specific and have not been identified in recent foodborne or waterborne outbreaks. The trend toward globalization of the food supply may also play a significant role in pathogen transmission. In Central America, 36% of irrigation water contained Cryptosporidium oocysts (ThurstonEnriquez et al., 2002), whereas in certain countries human sewage was used to fertilize horticultural crops. This high-risk practice leads to the spread of fecal pathogens to people who ingest such produce (Shuval, 1991). Untreated irrigation water is a significant concern because of the ubiquitous nature of Cryptosporidium, its low sedimentation rate, ability to retain its infectivity in moist environments (Rose, 1997), the presence of zoonotic species and genotypes, and the poor success of chemical treatments including chlorination (Smith and Nichols, 2006). There is a need to improve agricultural practices to reduce the burden of contamination. Restricting the access of cattle feces to watersheds and adequate treatment of feces for pathogen inactivation before disposal can reduce the transport of oocysts onto products that are consumed raw (Ong et al., 1996). Fertilization of horticultural crops only with adequately processed manure and use of clean water when spraying vegetables, either for irrigation or pesticide application purposes, will aid in reducing the burden of contamination of produce with viable Cryptosporidium as well as other pathogens. Additional control measures include education of producers and the development and implementation of effective measures to eliminate contamination of agricultural water and feed with viable stages of parasites (Gajadhar et al., 2006).
B.
Food Processing
Oocysts can also be introduced during food processing by direct contact with contaminated people, during the washing process, or by the addition of contaminated water as a significant ingredient in the foodstuff. This is particularly important in the preparation of beverages and juices. Oocysts can also be
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introduced when cleaning the processing or preparation equipment with contaminated water. The use of water sprinklers at the retail outlet display can also be a mechanism of spreading Cryptosporidium oocysts or other pathogens if the water is contaminated. Although outbreaks of cryptosporidiosis have not been associated with consumption of meats, the gastrointestinal contents of animals can be a source of infection for workers or surface contamination of meats during processing of carcasses in abattoirs.
C.
Pest Infestations
Pathogens may also be introduced by pest infestations such as cockroaches, house flies, mice, and rats. These can be reservoirs or mechanical vectors for Cryptosporidium because their excreta or body surface can harbor infectious oocysts. These pests can travel relatively long distances, transporting oocysts and depositing them in or near food. Because the infectious dose 50 of C. hominis can be as small as 10 oocysts, with a range of 10–83 oocysts in human volunteer studies (Chappell et al., 2006), pests may easily introduce this low number of parasites into foods.
D.
Food Handlers
Foods can also be contaminated by ill food handlers, as they can have symptomatic or asymptomatic infections. There are several documented reports that linked an infected handler with the occurrence of outbreaks (Table 10.2). Individuals generally excrete oocysts for 1–15 days after symptoms cease; however, it has been reported that oocyst excretion can last as long as 2 months (Jokipii and Jokipii, 1986). Thus, it is extremely important that good sanitation practices are implemented on a permanent basis among food handlers to prevent accidental introduction of Cryptosporidium and other pathogens into foods.
V.
Inactivation
Various mechanisms relevant to the agricultural and food industries have been evaluated for Cryptosporidium inactivation. The production of manure-based fertilizers should consider the following facts: (1) oocysts are very sensitive to desiccation and can be completely inactivated after 4 h in a dry environment at 18 to 20˚C or reach 95% inactivation at room temperature after 4 h (Deng and Cliver, 1999; Robertson et al., 1992); (2) oocysts can survive longer in slurries, feces, or moist conditions; (3) pH and organic content of diverse soil types may affect oocyst survival rates; and (4) oocysts can be inactivated when manure is composted and reaches temperatures exceeding 55˚C for 15 days (Van Herk et al., 2004). One study suggested that oocysts can survive at low pH solutions, although survival decreased with time (Friedman et al, 1997), whereas other studies provided evidence that pH may not significantly influence the viability of oocysts (Dawson et al, 2004). It has been reported that salinity reduces the oocyst survival; however, a small percentage of oocysts can remain viable for up to a year in seawater at 6 to 8˚C (Tamburrini and Pozio, 1999). It is also important to consider that because of the low specific gravity of the oocysts, sedimentation methods alone cannot be used to remove oocysts from water suspensions (Searcy et al., 2005). For beverages, hyperosmotic solutions (low water activity) have a detrimental effect on oocyst survival, similar to the presence of alcohol in certain beverages (Friedman et al., 1997). However, oocysts have been known to retain their viability in water, juice, and milk for long periods of time (Dawson et al., 2004; Deng and Cliver, 1999). High and low temperatures seem to be important determinants for oocyst survival. Freezing temperatures compromise oocyst survival, particularly when they are quickly frozen, but Cryptosporidium can survive if the temperature is slowly reduced. Therefore, oocysts that may be present in ice prepared with contaminated water may remain infectious (Olson et al., 2007). Additionally, oocysts may also retain their viability when frozen in an organic matrix.
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Oocysts can be removed from liquid matrices if filtered using 0.01 to 0.5 µm but the expense to implement this procedure may not be cost-effective. Similarly, irradiation, ultrasound, and high-pressure methods have been tested on Cryptosporidium oocysts (Ashokkumar et al., 2003; Peeters et al., 1989; Yu and Park, 2003), but these technologies have practical limitations. Ozone and UV light, effective for oocyst inactivation in water, had various levels of success with treatment of food, as it is highly dependent on time of exposure, dosage, oocyst concentration in the sample, and temperature (Korich et al., 1990). Therefore, careful validation studies are needed to ensure the efficacy of these methods when implemented in the production of food beverages.
VI. Hazard Analysis and Critical Control Points (HACCP) and Other Management Regulations Foodborne cryptosporidiosis could be drastically reduced if raw foods, other ingredients, and processing water were free of contamination. Thus, food processors must be very selective in the sources and suppliers of their raw food products. Product sampling and testing should be implemented to ensure that products meet the required microbiological standards. Stringency in the points of control will vary according to the products and processes. Food handlers can play a significant role in contamination. Educational programs that reinforce good sanitary practices and early detection of infected workers can play a significant role in preventing foodborne illnesses (Millard et al., 1994). Screening of ill food handlers must also consider that carriers or people who are in the convalescent stage of cryptosporidiosis may excrete oocysts for as long as 2 months after symptoms have stopped (Jokipii and Jokipii, 1986). Therefore, testing of stool samples from three consecutive days is important, as false negative results can be attributed to the excretion of low numbers of oocysts or intermittent excretion of parasites. Because water is a significant source of contamination, it should be parasite free when used to prepare ready-to-eat foods. The waterborne nature of Cryptosporidium was well documented in the 1993 Milwaukee outbreak in which more than 400,000 people acquired cryptosporidiosis by drinking contaminated tap water (MacKenzie et al., 1994). CFSAN released a guide to minimize microbial food safety hazards for fresh fruits and vegetables. It addressed the microbial food safety hazards and good agricultural and management practices for growing, harvesting, washing, sorting, packing, and transporting most fruits and vegetables sold and consumed in an unprocessed or minimally processed form. This guide was directed to domestic and foreign producers. Because agricultural practices and commodities are very diverse, the guide covers in a general fashion how each step could be adapted to specific operations to increase the safety of the produce (Anonymous, 1998b). Because of recent bacterial foodborne outbreaks, stakeholders, particularly the United Fresh Produce Association, will be releasing a document with goals for the fresh produce industry with specific recommendations to control fresh produce contamination on farms.
VII. Conclusions Food products have been associated with several outbreaks of cryptosporidiosis. Because most foodborne outbreaks are usually considered to be of bacterial or viral origin, the identification of Cryptosporidium or other parasites is often delayed, making epidemiological and trace-back studies more difficult. Therefore, the number of foodborne outbreaks of cryptosporidiosis is likely to be much higher than what is currently reported. Awareness in the health system along with better recovery and diagnostic assays for Cryptosporidium can facilitate outbreak investigations. There is a need to define multilevel strategies that involve food producers, processors, and retailers to prevent food contamination with Cryptosporidium. Additionally, if contamination occurred or was suspected to have occurred, there is a need to have an effective method
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to remove or inactivate the parasite in foods that is also compatible with food products, especially fresh produce, because conventional treatments including chlorination are not effective. The risk of acquiring cryptosporidiosis is not only associated with traveling to endemic areas where water or raw foods are more frequently contaminated with Cryptosporidium. The globalization of markets and trade may also affect the epidemiological patterns of infectious diseases. In the United States alone, the number of cases of cryptosporidiosis per million persons reported through FoodNet was 10.9 in 2003, 13.2 in 2004, and 29.5 in 2005 (Anonymous, 2004, 2005, 2006). Thus, there is a need to educate and implement preventive practices to minimize the burden of Cryptosporidium infections and to control factors associated with its transmission. This should not be limited to producers in industrialized nations but expanded to developing countries where fresh food products are grown or processed.
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11 Waterborne: Drinking Water
Jennifer L. Clancy and Thomas M. Hargy
CONTENTS I. II.
Introduction .................................................................................................................................. 305 Prevalence of Cryptosporidium in Water..................................................................................... 306 A. Prevalence in Early Studies ............................................................................................ 306 B. Prevalence in Recent Studies.......................................................................................... 308 III. Methods for Recovery of Cryptosporidium Oocysts in Water.................................................... 309 A. Sample Collection ........................................................................................................... 309 B. Sample Concentration ..................................................................................................... 311 C. Separation of Oocysts from Sample Debris ................................................................... 312 D. Detection in Water Samples............................................................................................ 313 E. Complete Methods .......................................................................................................... 314 Cryptosporidium-Monitoring Data Quality .................................................................... 315 F. IV. Drinking Water Regulations......................................................................................................... 316 A. U.K. Regulations............................................................................................................. 316 B. U.S. EPA Regulations ..................................................................................................... 317 V. Drinking Water Treatment............................................................................................................ 318 A. Filtration Technologies.................................................................................................... 318 1. Conventional Filtration .................................................................................... 318 2. Direct Filtration................................................................................................ 319 3. Slow Sand Filtration ........................................................................................ 319 4. Diatomaceous Earth Filtration......................................................................... 320 5. Bag or Cartridge Filtration .............................................................................. 320 6. Membrane Filtration ........................................................................................ 320 B. Disinfection ..................................................................................................................... 320 1. Ozone ............................................................................................................... 321 2. Chlorine Dioxide.............................................................................................. 321 3. Ultraviolet Light (UV) ..................................................................................... 322 VI. Conclusions .................................................................................................................................. 325 References........................................................................................................................... 326
I.
Introduction
Unlike waterborne diseases caused by bacteria such as cholera and typhoid, which have been controlled for over 100 years with the chlorination of water, the body of knowledge on effective treatment of Cryptosporidium in drinking water has been developing just over the past decade. Cryptosporidium parvum was recognized as an animal parasite over 90 years ago (Tyzzer, 1912), but it was not until the third largest waterborne outbreak in recent times that its importance as a significant human waterborne pathogen was recognized. The first known waterborne cryptosporidiosis outbreak with over 200 cases reported occurred in 1984 at Braun Station, TX, in a public water supply that was contaminated with
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sewage (D’Antonio et al., 1985). The water supply was groundwater receiving chlorination only. A second large outbreak affecting 6000 individuals occurred in the United Kingdom in 1987 in the areas of Oxford and Swindon, and was attributed to overwhelming levels of oocysts in the source water (Colbourne, 1989). However, it was the third outbreak in 1989 in Carrollton, GA, affecting about 13,000 people that began to change thinking in the water industry about Cryptosporidium and its role in waterborne disease (Hayes et al., 1989). Research on the parasite, including how to detect and control it in water supplies, was underway when the Milwaukee outbreak occurred in 1993, sickening 403,000 people and leading to over 100 deaths (MacKenzie et al., 1994). These large-scale waterborne disease outbreaks in the late 1980s and early 1990s, caused by a protozoan parasite that was virtually unknown to those outside the field of parasitology, shocked the drinking water community. Drinking water professionals thought the days of large waterborne disease outbreaks in the developed world had vanished with the widespread use of filtration and disinfection of drinking water supplies. The U.S. Environmental Protection Agency (USEPA) had finalized the Surface Water Treatment Rule (SWTR; USEPA, 1989) to control Giardia and viruses in drinking water. Cryptosporidium was not mentioned in the SWTR nor its companion publication, the Guidance Manual for Compliance with the Filtration and Disinfection Requirements for Public Water Systems Using Surface Water Sources (USEPA, 1990). In the wake of the outbreaks, the drinking water community responded quickly to develop methods for recovery and detection of oocysts in water to determine the occurrence, fate, and transport of Cryptosporidium in the environment. At the same time, studies were initiated to identify the capabilities of existing treatment processes for removing and/or inactivating Cryptosporidium, and to look for new approaches. As information on these areas of Cryptosporidium became available, regulatory agencies developed rules to control the parasite in water supplies to protect public health. This chapter describes the occurrence of Cryptosporidium in the aquatic environment, methods for recovery of oocysts in water, drinking water treatment for Cryptosporidium control, and regulation of Cryptosporidium in water supplies.
II. A.
Prevalence of Cryptosporidium in Water Prevalence in Early Studies
Cryptosporidium oocysts have been reported in both surface and ground waters, and in treated drinking water. The occurrence and levels of oocysts reported vary significantly, and are likely a factor of the methods used and expertise of the analysts. After the Carrollton outbreak, interest developed in the United States to learn more about the occurrence of Cryptosporidium in water. Madore et al. (1987) reported levels ranging from 0.8 to 5800 oocysts per liter in six samples in U.S. streams and rivers, with 100% of the samples testing positive. At that same time, Ongerth and Stibbs (1987) examined six rivers in California and Washington and found all 11 samples positive for oocysts, ranging from 2 to 112 oocysts per liter. The search for Cryptosporidium oocysts in water was underway. The first large-scale Cryptosporidium occurrence studies were reported in the early 1990s. Rose et al. (1991) examined surface water and groundwater used as drinking water sources in 17 states, categorizing sources as pristine (little or no human watershed activity, restricted watershed activity, no agricultural activity in the watershed, and no sewage discharges) or nonpristine. Maximum oocyst densities for nonpristine watersheds ranged from > 0.001 to 44 oocysts per liter, with oocysts in pristine watersheds detected in 39% of samples. One of 18 groundwater samples was positive at a concentration of 0.003 oocysts per liter. LeChevallier et al. (1991a) examined 66 source water supplies in 14 states and 1 province in Canada. Sources were characterized as protected from or impacted by industrial pollution. Of 85 samples collected, 87.1% were positive for oocysts, with concentrations ranging from 0.07 to 484/L. A similar study of finished drinking water indicated that oocyst levels in 82 samples ranged from 0.001 to 0.48/L, with 26.8% of samples positive (LeChevallier et al., 1991b). LeChevallier and Norton (1995) sampled previously studied sites and reported that 60.2% of 347 surface water samples were Cryptosporidium positive, with levels ranging from 0.065 to 65.1 oocysts per liter and a geometric mean
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of 2.4 oocysts per liter. In finished drinking water, 35 of 262 samples were positive, with oocyst levels ranging from 0.29 to 57 oocysts per 100 L, with an average of 3.3 per 100 L when detected. McTigue et al. (1998) conducted a year-long study of 100 surface water treatment plants in the United States, examining raw and treated drinking water. Cryptosporidium was detected in 77% of the raw water samples and in 15% of the treated water samples. In raw waters, oocyst levels ranged from 0.5 to 117 per 100 L and in finished drinking water the levels were 0.04 to 0.8 per 100 L. In a study of 463 groundwater samples from 199 sites in 23 of the contiguous 48 states, oocysts were found in 11% of the sites. Oocysts ranged from 0.2 to 45 per 100 L in the positive samples, averaging 5 per 100 L (Hancock et al., 1998). After the cryptosporidiosis outbreak in Oxford and Swindon (Colbourne, 1989), the Scottish Parasite Diagnostic Laboratory in Glasgow became very active in Cryptosporidium occurrence research. Smith et al. (1991) reported that 34 of 84 raw water samples contained oocysts at a concentration of 0.006 to 2.3/L. The National Cryptosporidium Survey Group (1992) examined source waters across the United Kingdom and reported lower Cryptosporidium levels than those initially reported in the early U.S. surveys. Ten sites of lowland waters had levels ranging from 0.04 to 4.0 oocysts per liter and < 1 to 57% of the samples were positive. The upper reaches of the systems had fewer oocysts than the urbanized sites. In Loch Lomond 11.5% of raw water samples were positive for Cryptosporidium (Parker et al., 1993). In central concentrations ranged from 0.0012 to 0.12 oocysts per liter (average = 0.02 oocysts per liter) in 15% of 403 raw water samples, whereas 1 of 15 finished water samples was positive at a concentration of 0.006/L (Humphreys et al., 1995). In Northern Ireland Lowery et al. (2001) conducted a multiyear study of Cryptosporidium in source waters and found 3% of 474 samples oocyst positive. Oocyst levels were generally quite low, ranging from 0.1 to 3.75/10 L; however, one source was highly contaminated during one sampling period, with oocyst levels from a feeder stream reaching 398 per 10 L. In the Greater Dublin area in Ireland, weekly water samples were collected from five sites (Skerrett and Holland, 2000). Of the 69 samples collected, 40.6% were positive for Cryptosporidium. Average numbers recovered at each site ranged from 0.3 to 1.1/L. In Germany, source waters at six water treatment plants the average number of oocysts was 116 per 100 L; in finished drinking water, 29.8% of 47 were positive for Cryptosporidium (Karanis et al., 1998). In Canada, pristine sites in the Yukon were found to be free of Cryptosporidium. Of the drinking water samples in Whitehorse 5% (2/42) were positive, and drinking water in Dawson was consistently negative (Roach et al., 1993). In British Columbia, two adjacent watersheds were sampled—the Black Mountain Irrigation District (BMID) and Vernon Irrigation District (VID)—that are unprotected and impacted by orchard farming and cattle ranching (Ong et al., 1996). At BMID, 52% of the intake samples were oocyst positive, ranging from 1.7 to 44.3 oocysts per liter. There was a statistically significant difference between up- and downstream oocyst levels, with spikes noted in downstream sampling. At VID, Cryptosporidium oocysts were detected in 29% of 19 samples over a 6-month period, with concentrations ranging from 4.8 to 51.4 oocysts per liter (geometric mean = 9.2 per 100 L). In samples from across Canada 4.5% of raw waters and 3.5% of treated waters were positive for Cryptosporidium, with positive results based on observing < 0.5 oocysts/100 L (Wallis et al., 1996). In the Montreal area, Payment and Franco (1993) found different concentrations of oocysts in the raw waters of three water treatment plants. Oocyst levels at the ROS plant ranged form 600 to 1200 per 100 L (geometric mean = 742 oocysts per 100 L) with 100% of samples positive; the finished drinking water had 1 of 9 positive samples with a geometric mean of 0.02 oocysts per 100 L. At the STE plant, only 1 of 8 raw water samples were positive at a geometric mean level of < 2 oocysts per 100 L (range = > 2 to 20 oocysts per 100 L). At the REP plant, none of the raw (0 of 5) or finished (0 of 13) samples were positive (< 2 per 100 L), but oocysts were detected in the settled water samples. In Ottawa, Ontario, Chauret et al. (1995) examined the levels of Cryptosporidium oocysts in two rivers, finding levels as high as 2 oocysts per liter in the Ottawa River and 100 oocysts per liter in the Rideau River. The first studies of Cryptosporidium in Australia were conducted for the Sydney Water Board by Hutton et al. (1995). In raw water storage reservoirs, levels ranged from < 0.3 to 42.9 oocysts per liter. In the Nepean River and Warragamba Dam, the average oocyst densities were 0.87 oocysts per liter and 0.69 oocysts per liter, respectively; in the distribution system Cryptosporidium was detected at a level of 0.21 oocysts per liter. In rural Australia, Thurman et al. (1998) studied water quality in four creeks
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and two drinking water reservoirs. The levels reported were low, ranging from 0 to 6 oocysts per 10 L in the source waters, with averages ranging from 0.38 to 2.67 per 10 L at the four creek locations, and 4 per 10 L at both reservoirs. In Asia, Hsu et al. (1999) found high levels of oocysts in the watershed area and the Kau-Ping River, which serves 2.5 million people in Taiwan. Raw water samples (75% positive) were as high as 12,516 oocysts per 100 L at one site, whereas treated drinking water levels (40% of samples positive) were reported as high as 159 oocysts per 100 L. In Hong Kong, in the Lam Tsuen River, oocysts were found at one of four sampling sites each at concentrations of 3 to 30 per 10 L; along the Shing Mun River, a single site was positive for Cryptosporidium once with 13 oocysts per 10 L (Ho and Tan, 1998). In Israel, 80% of 15 samples from five streams were positive for Cryptosporidium with an average concentration of 0.04 to 1.9 oocysts per liter (Zuckerman et al, 1997). Two springs were positive for Cryptosporidium at a concentration of 0.54 oocysts per liter. Cryptosporidium was isolated in 4 of 6 samples at an average concentration of 0.3 to 1.09 oocysts per liter from Lake Kineret, the main drinking water reservoir of Israel. Drinking water entering a filtration pilot plant was positive for Cryptosporidium in 23 of 35 samples (0 to 317 oocysts per liter).
B.
Prevalence in Recent Studies
The first required monitoring for Cryptosporidium was in the United States, initiated by the U.S. EPA under the Information Collection Rule or ICR (USEPA, 1997). The ICR required utilities serving more than 100,000 people and using surface water to monitor source waters for Giardia cysts, Cryptosporidium oocysts, and culturable viruses for 18 months. Source waters found to be positive for either protozoan at greater than or equal to 1000 per 100 L, or positive for culturable enteric viruses greater than or equal to 100 per 100 L, required finished water monitoring. A quality assurance program was instituted and specific methods, laboratory audits, and proficiency testing for protozoan and enteric virus testing were prescribed by the USEPA (USEPA, 1996). Unlike the majority of the previously discussed studies, 93% of the 5829 samples analyzed for the protozoa did not detect Cryptosporidium. Significant method improvements for Cryptosporidium were underway in 1997, but not in time for the ICR monitoring which was legislated using the ICR method. The USEPA published the first version of its new method—Method 1622—in 1998, and many laboratories began to use it while it was being refined (USEPA, 1998). Development of Method 1622 is discussed in Section III E. In a prefecture in western Japan all large rivers used as sources of water supply were examined by improved methods (Ono et al., 2001). One sample was collected at each of 156 sites along 18 rivers. Samples were classified as obtained from (1) an island with livestock and fishing industries, (2) a densely populated urban area, (3) a western region including farming villages, or (4) a more rural northern area with agriculture and fishing. Cryptosporidium was detected in at least one sample from 72% of the 18 rivers and in 47% of 156 samples. One-third to all of the samples from each area contained C. parvum oocysts. In the positive samples, there were 1 to 8 Cryptosporidium oocysts per 20 L of river water, with an average of 1.8 per 20 L. Genotyping of three water samples showed the presence of C. parvum. A Japanese water purification plant monitored for Cryptosporidium oocysts for one year (Hashimoto et al., 2002). Thirteen 50-L samples of river source water and 26 samples of 2000-L filtered water, treated by coagulation, flocculation, sedimentation, and rapid filtration, were tested. Cryptosporidium oocysts were detected in all 13 raw water samples, with a mean concentration of 40 oocysts per 100 L. In the filtered water samples, oocysts were detected in 35% of the 26 samples with a geometric mean concentration of 1.2 oocysts per 1000 L. In Finland a year-long study used improved methods to examine waterborne pathogens from seven lakes and 15 rivers in the southwest, looking for correlations between pathogen and indicator detection (Hörman et al., 2004). Levels of oocysts were not reported, but Cryptosporidium spp. were isolated from 10.1% of 139 samples. In North America an intensive study of six watersheds, including two runoff seasons, was conducted to examine pathogen levels in source waters in very different watersheds (LeChevallier et al., 2003). Data are shown in Table 11.1. The 593 samples were analyzed using Method 1622, by a person with over 10 years’ experience in water analysis of Cryptosporidium, and recoveries in matrix spike samples
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TABLE 11.1 Levels of Cryptosporidium in North American Watersheds Watershed
Characterization
Number of Samples
Average/L
Range/L
Bull Run Reservoir, OR Allatoona Lake, GA Mianus River, CT Mississippi River, IO Tennessee River, TN Grand River, ON
Fully protected Recreation, sewage, agriculture, CSO Limited access, recreation, agriculture Recreation, agriculture, sewage, CSO, industry Agriculture, sewage, CSO, industry Recreation, agriculture, sewage, industry
97 98 98 101 101 98
0.004 < 0.001 0.012 0.017 0.005 0.069
0.0–0.08 — 0.0–0.50 0.0–1.17 0.0–0.20 0.0–1.00
Note: CSO = Combined sewer overflow. Source: From LeChevallier, M.W., DiGiovanni, G.D., Clancy, J.L., Bukhari, Z., Bukhari, S., Rosen, J.S., Sobrinho, J. and Frey, M.M. 2003. Comparison of Method 1623 and cell culture-PCR for detection of Cryptosporidium in source waters. Appl. Environ. Microbiol. 69, 971–979. With permission.
exceeded 70%, providing excellent quality control and quality assurance. The most agriculturally impacted watershed was the Grand River in Ontario, where the concentration of oocysts averaged 0.069/L, but did not exceed 1/L. The most pristine watershed, the Bull Run Reservoir serving Portland, OR, provided disinfection only. Characterized as fully protected, it had an average concentration of 0.004 oocysts per liter and did not exceed 0.08 oocysts per liter. In the province of Álava, northern Spain, a 30-month study of Cryptosporidium examined 284 samples from drinking and recreational water supplies (Carmena et al., 2007). Oocysts were found in 63.5% of river samples; 33.3% of reservoirs samples; 15.4 and 22.6% of raw water samples from conventional and small water treatment facilities, respectively; 30.8% of treated water from small treatment facilities; and 26.8% of tap water from municipalities with chlorination treatment only. Natural surface water from rivers and reservoirs had the highest occurrence of protozoa, with concentrations that reached 1767 oocysts per 100 L. Both finished drinking water (26%) and tap water samples (31%) contained levels of < 8 oocysts per 100 L. Cryptosporidium monitoring continues worldwide, with the majority of monitoring done by utilities on an intermittent basis to develop internal data. The United Kingdom has a required finished drinking water monitoring scheme for utilities at risk for Cryptosporidium, and the United States has recently instituted a source water monitoring program. Both are described in Section IV.
III. Methods for Recovery of Cryptosporidium Oocysts in Water Cryptosporidium analytical methods for water are characterized by four components: sample collection, concentration, separation of oocysts from sample debris, and detection. Since the first isolation of oocysts from water, a variety of techniques have been examined for each method component. This section will review the techniques that have been used for Cryptosporidium analysis of water, the history of method development and validation, and describe the current methods that are used commonly.
A.
Sample Collection
Filters used for Cryptosporidium sampling are generally used for Giardia sampling as well, because both waterborne parasites are of interest to water suppliers and public health officials. In 1979, the USEPA developed the first practical sample collection method for Giardia, which was a portable system that consisted of a nominal porosity string wound filter for sample collection (Jakubowski and Erickson, 1979). The process involved washing the filter with distilled water to remove particulates for further processing and microscopic examination. This method was not used routinely but was developed for investigation of waterborne disease outbreaks. When interest in Cryptosporidium developed, the method was used to test for oocysts.
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Two methods for Cryptosporidium recovery and identification were published in 1987. A complete method using the 1-µm nominal pore size polypropylene wound cartridge filter rather than the string filter used earlier was developed by Musial et al. (1987). The 1 µm nominal pore size string or polypropylene wound cartridge filter was the predominant type of filter used for sample collection for over 10 years. It is still used in some countries because it is very inexpensive. An advantage is that very large volumes of water, even turbid water, can be passed through the filter; the drawback is that a nominal pore size permits passage of particles much larger than the nominal rating to pass through the filter. Greater than 60% of oocysts were shown to pass through nominal-pore-size filters (Clancy, 1996). A polycarbonate-track-etched (PCTE) flat membrane with a precise pore size of 5 µm was developed by Ongerth and Stibbs (1987). The water sample filtrate from the 5 µm filter was collected and refiltered through a PCTE 1 µm pore size membrane and further processed. This process was later changed to omit the prefiltration step. A 1 or 2 µm PCTE membrane was used for direct sampling, the membrane was washed, and the rinse material was collected (Nieminski et al., 1995). The drawback of this filter is that it is limited to small sample volumes because the pores clog quickly with raw water and with certain types of treated drinking water; a significant advantage is that none of the particles of interest pass through the filter (Clancy et al., 1997). A comparison of the wound cartridge and membrane filter methods in Taiwan found the membrane yielded higher recovery and detection rates than the wound filter (Hsu et al., 2001). Another method using a flat membrane made of cellulose acetate to filter the water, followed by dissolution of the membrane in acetone, was developed in a Canadian laboratory (Aldom and Chagla, 1995). This method had the advantage of preventing potential loss of oocysts in the filter-rinsing steps. Oocyst recoveries using 1.2- and 8.0-µm pore-size membranes were 70.5 and 56%, respectively (Aldom and Chagla, 1997). Graczyk et al. (1997) obtained 78.8% oocyst recovery using this method, but McCuin et al. (2000) conducted studies with low seed doses of oocysts and found recoveries to be low and variable. The membrane dissolution method was popular in Canada and Japan for Cryptosporidium sample collection. Wound fiberglass depth cartridge filters used in Australia obtained oocyst recoveries of 43 to 84%, averaging 54% (Kaucner and Stinear, 1998). This method was based upon a method of Payment et al. (1989) used to recover multiple pathogens and indicators from drinking water. Cartridge filtration, membrane filtration with three types of membrane materials, and calcium carbonate flocculation (described in section IIIB) were examined by Shepherd and Wyn-Jones (1996) who found that cellulose acetate membranes with a 1.2 µm pore size gave the best oocyst recovery. A study of ultrafiltration using a polysulfone hollow fiber single-use filter showed Cryptosporidium recoveries of 42% in both method blank and raw water samples (Simmons et al., 2001). A hollow-fiber ultrafiltration system (50,000 MWCO) was optimized by Kuhn and Oshima (2001, 2002) to concentrate oocysts from 10 L samples of environmental water. Its performance was similar to the Envirochek capsule in waters of low turbidity waters (~3.9 ntu), with higher recoveries in highly turbid samples (159 ntu), but cautioned that sample numbers were limited. A novel filter system using open cell reticulated foam rings compressed between retaining plates and fitted into a filtration housing was evaluated in the United Kingdom for the recovery of oocysts from water. Mean recoveries of 90.2% from seeded small and large volume (100 to 2000 L) tap water samples, and 88.8% from 10 to 20 L river water samples, were achieved. This was a marked improvement in capture and recovery of Cryptosporidium oocysts from water compared with conventional polypropylene wound cartridge filters and membrane filters (Sartory et al.,1998). This filter, the IDEXX Filta-Max® is approved for Cryptosporidium monitoring in the United Kingdom and United States; a similar new product, the Filta-Max xpress is now approved in the United States. Laboratories in the United States and United Kingdom were very active in the development of Cryptosporidium analytical methods, and a joint U.S.–U.K. effort led to the development and testing of the Pall Envirochek® capsule. Water samples are filtered through a Supor® polyethersulphone membrane with a 1-µm absolute pore size, allowing complete capture. Oocysts are mechanically eluted from the membrane fiber using a wrist action shaker and a nonionic detergent. This was the first method to allow multiple samples to be processed within 1 h. Recoveries of C. parvum oocysts from seeded tap and source water samples were 90 to 95% (Matheson et al., 1998). This filter is approved for Cryptosporidium
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monitoring in the United Kingdom and United States. The Envirochek HV was developed for highvolume sampling and can be used for both source and treated drinking water. A third filter, the Whatman CrypTest®, was approved by the USEPA for Cryptosporidium testing (Clancy et al., 2003). This is a closed cartridge filter of pleated PCTE membrane in a reusable holder.
B.
Sample Concentration
Once a water sample is collected on a filter, the collected material has to be washed from the filter and concentrated into a small volume that will eventually end up on a microscope slide for identification and enumeration of Cryptosporidium oocysts. For wound cartridge filters, a scalpel is used to cut the filter fibers to the plastic core; the fibers are pulled apart and hand-washed in buffer with or without detergent. The introduction of the Stomacher® was an improvement. The Stomacher® is a mechanical device into which a disposable bag with detergents and filter fibers are placed and a mechanical arm provides the washing. This replaced hand washing in most laboratories, and added standardization to the wash time and possibly the effectiveness of the procedure of particulate removal. For the PCTE membrane filters, the membrane was inverted in distilled water (DI) and vibrated to remove the particulates (Ongerth and Stibbs, 1987). Later adaptations included scraping the membrane or handwashing the membrane in DI or buffer and detergents. For the cellulose acetate membranes, particulate concentration was done when the membrane was dissolved in acetone (Aldom and Chagla, 1995, 1997). Several products use a washing technique to remove the sample from the filter. The Envirochek® capsule is a single use, disposable unit and is eluted by mechanical shaking with a Laureth12 buffer to remove oocysts and other particulates from the pleated membrane (Matheson et al., 1998; Clancy et al., 1999). The CrypTest® cartridge filter is backwashed in the filter holder using an air purge and elution buffer; the cartridge is a single use but the holder is reusable. The Filta-Max is eluted by opening the holder and releasing the foam pads, followed by washing them in buffer using a manual or automatic wash station. A new product called the Filta-Max xpress® uses a fully automated elution station where an air pressure backwash forces the elution buffer through the filter. Gravity settling was the earliest method used for concentration. Filter fibers were hand-washed in an elution solution, wrung to remove as much of the liquid as possible, and fibers discarded. The liquid, usually several liters, was allowed to settle in a large beaker overnight to collect particulates hand-washed from the filter fibers. The next day the liquid potion was removed, and the resulting sediment collected and further processed. Slow-speed centrifugation was an alternate method for concentration. Samples from filter washings were poured into centrifuge bottles and centrifuges at any of a variety of speeds, often dependent on the type of centrifuge available in the laboratory. Variations in centrifugation speeds, bottle volume and design, rotor types, refrigerated or not, and rotor braking or not have been used (Musial et al., 1987; Ongerth and Stibbs, 1987; Anon., 1991; LeChevallier et al., 1990, 1995; Clancy et al., 2000a). Standard methods have specifications for sample concentration using centrifugation, but these vary among methods. Most methods currently use low-speed centrifugation (1500 to 4500 × g), plastic conical bottles ranging from 250 to 500 mL, and swinging bucket rotors with no brake in unrefrigerated centrifuges for sample concentration. After centrifugation, the supernatant fluid is poured or aspirated from the bottle and the resulting material, referred to as the packed pellet, subjected to further processing. However, four techniques have been developed that combine both the sample collection and concentration into a single step. For samples where turbidity is high and only a small volume could be filtered, it is acceptable to perform direct centrifugation of the sample itself without prior processing. A grab sample is collected, poured into centrifuge bottles, and centrifuged using some combination of parameters as described previously. The supernatant fluid is removed and the resulting packed pellet subject to further processing in the same manner as a sample that was filtered and eluted prior to centrifugation. A calcium carbonate flocculation method for oocyst recovery from water was developed in the United Kingdom. Calcium chloride is added to a 10-L grab sample and mixed well, followed by the addition of sodium carbonate. Oocysts are concentrated in the formation of calcium carbonate and released from
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this matrix with the addition of sulfamic acid, with recoveries of ~68% (Vesey et al., 1993a). This method became the standard in the United Kingdom, whereas in the United States the polypropylene wound cartridge filter remained the standard. Nearly a decade later, Karanis and Kimura (2002) conducted a comparison of three flocculation methods using ferric sulfate, aluminum sulfate, and calcium carbonate; the indication was that twice as many oocysts were recovered with ferric sulfate (61.5%) than with calcium carbonate (38.3%). However, by this time few labs were using flocculation for sample concentration. Vortex flow filtration (VFF) was first used by Whitmore and Carrington (1993) for the concentration of oocysts from water. Fricker and coworkers (Fricker et al., 1995, 1997) achieved recoveries of 70% consistently using VFF for concentration of protozoa from all types of water samples. Clancy et al. (1997) tested VFF as a possible replacement for the polypropylene wound cartridge in the United States, but a long operator learning curve and problems handling water qualities in excess of 5 ntu were noted; hence, it was not recommended. A stationary differential continuous flow centrifuge usually employed for blood cell separation was developed it into a device for concentrating oocysts from large volumes of water (Zuckerman et al., 1999). Oocysts are compacted onto the wall of a disposable plastic centrifuge bowl, residual water is aspirated, and the oocysts dislodged by detergents and vigorous shaking. Oocyst recoveries were about 90%. This method was approved by the U.S. EPA for Cryptosporidium monitoring (Zuckerman and Tzipori, 2006). The continuous flow (CF) centrifuge was evaluated for recovering oocysts from water under realistic environmental conditions (Swales and Wright, 2000). The maximum percent recoveries were achieved at a flow rate of 0.75 L/min and centrifuge speed of 2900 × g. When compared directly with cartridge filtration at a seeding level of 10 oocysts per liter and low turbidity of 1 ntu, recoveries were 14.8 and 9.7% for the CF centrifuge and cartridge filtration, respectively. At 5 ntu, recoveries were 13.0 and 9.0%, respectively. They concluded that these differences were not significant but the CF centrifuge sample was faster, simpler, and safer to process. Borchardt and Spencer (2002) investigated continuous separation channel centrifugation for Cryptosporidium oocyst recovery from water, with similar results. In tap water samples, C. parvum oocysts recovery ranged between 37.4 and 84% with a trend for higher recovery with higher oocysts densities, averaging 66.9%, whereas 1000-L tap water samples showed a mean recovery of 35%. Higgins et al. (2003) found that continuous flow centrifugation was not as efficient as capsule-filtration based methods, but that CFC and IMS could provide a more rapid and economical alternative for isolation of C. parvum oocysts from highly turbid water samples containing small quantities of oocysts.
C.
Separation of Oocysts from Sample Debris
For oocysts to be identified in a water sample they must be separated from debris that results from sample concentration. The earliest environmental methods relied on buoyant density gradient flotation for separation. Generally, the packed pellet is layered on the density gradient and centrifuged; biological particles of interest, including oocysts, are harvested at and just above the interface level. This floated suspension is rinsed and further concentrated by centrifugation prior to analysis. The highest oocyst recoveries were obtained when the packed pellet material was treated with 1% Tween 80 and 1% SDS and sonicated prior to flotation on Sheather’s sugar solution (Musial et al., 1987). An alternate method was developed about that same time using 40% potassium citrate (s.g., 1.195) for oocyst separation (Ongerth and Stibbs, 1987). In 1990, a method was introduced using a Percoll (polyvinyl pyrrolidone-coated silica particles)–sucrose flotation (LeChevallier et al., 1990). A 1.0-mL volume of packed pellet was adjusted to 20 mL, underlayered with 30 mL of Percoll-sucrose flotation medium (sp. gr., 1.09) and centrifuged at 800 × g for 5 min. The top water layer was drawn off and further processed. Flotation was better than trying to examine samples directly as packed pellet material, but the results with flotation were highly variable due to loss of oocysts with the pellet debris and flotation of other biological particulates such as algae that obscured oocyst detection. A flow cytometry method was developed for separation of oocysts from other sample debris (Vesey et al., 1993b). A flow cytometer is a laser instrument that can be equipped with fluorescence activated
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cell sorting (FACS), allowing it to analyze cells as they travel in a moving fluid stream (known as the sheath fluid) past the fixed laser beam. As oocysts, stained with a fluorochrome dye, pass in front of the laser, several measurements are made based on the physical characteristics of the cell. These characteristics, which pertain to how the cell scatters the laser light and emits fluorescence, will provide information about the cell’s size, internal complexity, and relative fluorescent intensity. The FACS sorts these cells onto a slide for microscopic confirmation. Flow cytometry for separation of oocysts from water samples became the method of choice in the United Kingdom and Australia for several years (Veal et al., 1997). Development of the first kit for immunomagnetic separation (IMS) of Cryptosporidium was a significant method improvement (Campbell et al., 1997). IMS uses paramagnetic beads coated with Cryptosporidium antibody to bind to antigens on the oocyst surface. The beads and oocysts from the sample matrix form an oocyst–bead complex. The concentration process is created by a magnet placed on the side of the test tube bringing the oocyst–bead complex to the magnet. The IMS buffer is removed from the sample and the bead–oocyst complex dissociated; the beads are removed from the sample using the magnet, leaving the purified oocysts for detection. Although flow cytometry was a significant improvement over buoyant density gradient flotation, the instruments are expensive to buy and maintain, and operator experience is critical. IMS is a simple technique requiring minimal investment in equipment and accessible to routine monitoring laboratories. Studies have shown that IMS permits recovery of oocysts from highly turbid samples (Bukhari et al., 1998) and does not affect oocyst infectivity (Rochelle et al., 1998). IMS is the most commonly used method for oocyst separation from water sample debris.
D.
Detection in Water Samples
Fluorescent monoclonal antibodies (mAb) for Cryptosporidium oocysts were developed for clinical use (Sterling and Arrowood, 1986; Arrowood and Sterling, 1989), but oocyst detection in water samples would not be possible without this important tool (Stetzenbach et al., 1988; Rose et al., 1989). The small oocyst size, coupled with other particles of similar size and shape in water, make oocyst identification very difficult or impossible. Early methods for Cryptosporidium detection in water employed sample concentration by filtration, separation using density gradients, and monoclonal antibody staining, but were referred to for years as the immunofluorescence or IFA method. Oocysts can be stained with fluorescent mAbs in solution, after drying on microscopic slides, or on membranes. The mAbs are generally conjugated to fluorescein isothiocynate (FITC), but other fluorochromes can be used. Currently, most methods use IFA staining after drying the oocyst preparations on slides. Hoffman et al. (1999) compared four commercially available FITC-labeled mAb kits for avidity, lot-to-lot variability, and crossreactivity, finding that the kits differed in each of these aspects. No kit demonstrated excellent performance overall. Cross-reactivity even using mAbs in environmental samples is an issue; algae and other organisms common in water samples react with the mAbs, either through antigen recognition or nonspecific binding. A similar study was undertaken by Ferrari et al (1999) and found that mAb of the IgG1 subclass to Cryptosporidium resulted in a more reduced background fluorescence than those prepared with the IgM or IgG3 subclass. When examining water samples for the presence of Cryptosporidium oocysts, the fluorescence pattern and the size and shape of the object are used as the initial discriminators. Slides are scanned at 200× magnification and if the object of interest is 4 to 6 µm in diameter, round, with a brightly fluorescing outer rim, the object is then examined at 400×; if it still fits the morphological criteria, it is examined at 1000×, generally followed by some method of confirmation, including DIC microscopy and inclusion of DAPI-stained sporozoite nuclei (Grimason et al., 1994). This is the most commonly used method for oocyst detection and enumeration in water samples. Laser scanning cytometry detects, enumerates, and records the presence of mAb-labeled Cryptosporidium oocysts on microscope slides (Reynolds et al.,1999; de Roubin et al., 2002). The laser scanning apparatus is a laser scanning unit equipped with a 488-nm argon laser that scans a 9-mm glass single well microscope slide in 3 min. Photomultiplier tubes detect the fluorescence light emitted by the labeled oocysts. The signals produced are processed by a computer using a series of proprietary software discriminants that enable the instrument to differentiate between valid signals (labeled oocysts) and
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background noise (e.g., autofluorescent particles and debris). Laser scanning results are displayed on a scan map and are visually confirmed using epifluorescence and DIC microscopy. This method reduces time, operator fatigue, and subjectivity in scanning slides where workloads are high. This method has been approved in the United Kingdom for Cryptosporidium monitoring. Other methods have been developed for detection of oocysts including flow cytometry (Vesey et al.,1994), spectroscopy (Callahan et al., 2003), immunoassays (Siddon et al., 1992), probe hybridization (Aguilar and Fritsch, 2003), in situ hybridization (Smith et al., 2004), Nucleic Acid Sequence Based Amplification (NASBA) (Kozwich et al., 2000), polymerase chain reaction (PCR) (Johnson et al., 1993), reverse transcriptase PCR (RT-PCR) (Rochelle et al., 1997a; Kaucner and Stinear (1998), nested PCR (Monis and Saint, 2001), and real-time PCR (Guy et al., 2003). None of these methods have been developed for routine use in Cryptosporidium monitoring, and they remain tools of research laboratories. Development of C. parvum in cell culture was first reported in 1984 (Current and Haynes, 1984) and in the 1990s many cell lines were demonstrated to support infection. Cell-culture-based infectivity assays were developed specifically for drinking water applications (Rochelle et al., 1997b; Slifko et al, 1997; DiGiovanni et al., 1999) and the equivalency of cell culture with a standard mouse infectivity assay has been demonstrated (Rochelle et al., 2002). A study utilizing cell cultures to assess infectivity reported that 27% of surface water treatment plants were releasing infectious oocysts in their finished water and, overall, 1.4% of treated drinking water samples contained infectious oocysts (Aboytes et al., 2004). Although cell culture is not used routinely, it is used in research studies to assist in determining whether the oocysts detected in water or wastewater effluents are viable and present a health risk.
E.
Complete Methods
There are dozens of ways that each of the previously discussed techniques can be organized into complete methods for Cryptosporidium oocyst recovery, separation, detection, and enumeration in water. In some countries, methods are developed and recommended for monitoring, but only when the monitoring is regulated by a health authority is there scrutiny in all aspects of method development. Cryptosporidium monitoring is currently required in the United States and United Kingdom, and although the regulatory agencies in each country approach this monitoring water quite differently, the methods used are very similar (Clancy and Hunter, 2004). When monitoring becomes required, there is a legal basis for gathering data, and as part of this scheme, a complete method must be developed and tested. As such, the methods for monitoring in the United States and United Kingdom are similar in that they prescribe specific methods, require methods approval and validation, use a laboratory approval process, and conduct ongoing proficiency testing for quality assurance/quality control (QA/QC). Although variations of the techniques are used worldwide, the majority of the testing relies on the Method 1622, which is very similar to the U.K. method—Standard Operating Protocol for the Monitoring of Cryptosporidium Oocysts in Treated Water Supplies to Satisfy the Water Supply (Water Quality Regulations) in England and Wales—used for treated water monitoring (http://www.dwi.gov.uk/regs/crypto). Method 1622 was developed as an improvement to the ICR method shortly after the ICR monitoring had begun. The problems with the ICR protozoan monitoring method are well documented and include poor reproducibility and sensitivity, high detection limit (>100 organisms per liter), the inability to differentiate Cryptosporidium using IFA-based technology, and both a high false-positive and falsenegative rate. The method is technically difficult, and a 16-laboratory collaborative study of the ICR method showed that overall performance was poor in laboratories that routinely used this method (Clancy et al., 1994). Attempts at improving the method were not successful (LeChevallier et al., 1990, 1991; Clancy et al., 1997). Method 1622 was developed through the U.S. EPA’s Office of Science and Technology to provide a reliable method for Cryptosporidium in water. The approach taken to developing Method 1622 was to evaluate each step in the method—sampling, processing, and assay—for its ability to permit recovery of spiked oocysts in reagent water. Each individual step was optimized in reagent water, and after optimization, the steps were combined into a full method and validated in two laboratories (Clancy et al., 1999). The method uses specified filtration techniques, separation of oocysts from sample debris using IMS, drying the sample on a microscope slide and staining with FITC-labeled mAbs and 4-6-diamidino-2-phenylindole (DAPI), and observing the oocysts at 1000× magnification under DIC
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(USEPA, 2001). Originally developed for 10-L source water samples, the methods have been adapted for higher volume source (50 L) and finished water (1000 L) monitoring (McCuin and Clancy, 2003). Method 1622, Cryptosporidium in water by filtration/IMS/FA, can be found at www.epa.gov/microbes. Method 1622 is still characterized by high variability, but is much more sensitive than previous methods. In laboratories with well-trained analysts, recoveries can be 50% or greater routinely with a seed dose of 100 organisms. With the improvements in sampling, separation, and staining, false-positive and falsenegative rates are lower. These methods are significant improvements over the ICR and other methods, and recognizing this, many countries have adopted them for use as the standard for Cryptosporidium analysis of water.
F.
Cryptosporidium-Monitoring Data Quality
Monitoring data on Cryptosporidium occurrence in water have been gathered since the late 1980s by researchers worldwide using a variety of techniques and methods. Even after the development of improved techniques, some laboratories continue to use the older methods including, the string- or propylene-wound filters, usually due to the much higher cost of the improved filters; flotation rather than IMS, again, often due to the cost; and detection with IFA only, rather than with DAPI and DIC to confirm oocysts. When examining occurrence data, it is important to note the methods used and judge the reliability of the data accordingly. Analysis of the 18 months of ICR data showed that of 5,829 samples analyzed for the protozoa, 93% of the samples were nondetects for Cryptosporidium. When oocysts were detected, it was generally one or two organisms observed in a small subsample. These observed numbers were then extrapolated to 100 L for source water or 1000 L for treated water. The final result was often a very high reported number, based on this extrapolation. The ICR data fall generally into two broad categories—either nondetects or very high reported levels based on low analyzed sample volume. For example, from a 100-L sample, a portion equivalent to ~2.5 L is actually examined microscopically due to method limitations. If one oocyst is observed, then the reported value is 40 oocysts per 100 L, assuming incorrectly that oocysts are evenly distributed in a sample. For nondetect data, if no oocysts were detected in the 2.5-L equivalent volume, the count is reported as < 40 oocysts per 100 L. This means the count could be 39 or 0 or any number in between. The method is so poor that actual count data as well as nondetect data are unreliable (Allen et al., 2000). A number of papers have argued for reporting the actual oocyst count per volume examined and avoiding extrapolation (Parkhurst and Stein, 1998; Atherholt and Korn, 1999; Clancy et al., 1999; Young and Komisar, 1999). Reporting of actual oocyst counts per volume of sampled examined is a hallmark of Method 1622. A majority of the studies discussed earlier used the early protozoan methods, which lacked both sensitivity and specificity; these methods include the ICR method and its forerunners, developed in the United States; the Standing Committee of Analysts method (U.K.); and variations on these methods. One notable difference after the ICR method was introduced was the use of DIC of phase-contrast optics to examine potential oocyst-like objects. The ICR data stand in stark contrast to the high levels noted in the early monitoring studies. One possible explanation is that the early versions of the IFA method did not require confirmation of oocysts, but relied on IFA identification alone, and so overestimates (false-positives) were likely to be reported. Reliable Cryptosporidium occurrence data depend on the methods and expertise of the analysts. Even with improved methods for both capture and recovery of oocysts, the microscopic identification remains subjective and correct identification depends on the skill of the microscopist (Clancy, 2000b). When reviewing Cryptosporidium occurrence data, a careful examination of the methods used and the experience of the analysts can provide insight into data reliability. Methods for recovery of Cryptosporidium in water have been improved, but limitations remain. Method 1622 has high variability depending on water quality and laboratory expertise, and there is no way to determine oocyst species or if the oocysts detected are viable or infectious to humans. Weintraub (2006) has suggested that better Cryptosporidium testing methods will improve confidence in the data and will simplify the risk communication task, and the level of trust which the public has in the water utility can be maintained and improved. Although improved analytical methods are needed to understand the
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occurrence, fate, and transport, species, and infectivity of Cryptosporidium oocysts, testing cannot be relied on for predicting or preventing disease.
IV.
Drinking Water Regulations
In the last 15 years, our understanding of the occurrence of Cryptosporidium in water and the effectiveness of water treatment have improved significantly. With this information, regulatory agencies have developed regulations to protect public health by controlling the parasite through monitoring, removal, or inactivation. Two different approaches have been developed in the United States and United Kingdom (Clancy and Hunter, 2004).
A.
U.K. Regulations
After a number of large waterborne disease outbreaks, the United Kingdom was first to develop specific regulations to control Cryptosporidium. The U.K. Drinking Water Inspectorate requires treatment to achieve a minimal concentration of Cryptosporidium in finished drinking water verified by full-time monitoring (Water Supply Regulations, 1999; 2000), and does not distinguish between infectious, noninfectious, or dead oocysts. As a result, removal (filtration) processes are favored, whereas disinfection processes, which do not lower oocyst concentration, confer no advantage toward meeting minimum concentrations as regulations demand. The regulations require that water intended for human consumption “is free from any microorganisms and parasites and from any substance which, in numbers or concentrations, constitute a danger to human health.” The regulations for testing water supplies for Cryptosporidium were originally set out in the Water Supply (Water Quality) (Amendment) Regulations (1999). These regulations were then incorporated into the Water Supply (Water Quality) Regulations (2000). These regulations require water utilities to undertake a number of steps. The first of these is to conduct a risk assessment to determine whether each supply is at significant risk from Cryptosporidium. Supplies considered at significant risk include: • • •
Those using direct withdrawal or with an average storage of 7 days or less from a river or stream. Those showing evidence of rapid river or surface water connection to the aquifer demonstrated by the confirmed presence of fecal coliform bacteria in the raw water. Those with a past history of an outbreak of cryptosporidiosis associated with the water supply where the reason is unexplained and no specific steps have been taken to prevent a recurrence.
If a water supply is deemed to be at increased risk of Cryptosporidium oocysts, then the water supplier is required to put in place a water treatment process that will guarantee an oocyst count of <1/10 L and have monitoring in place to demonstrate that this standard is being met. This standard is not health- or risk-based, but is considered to be achievable in a properly operated treatment plant. The sampling process is continuous and at least 40 L/h needs to be filtered. A gap in monitoring of no more than 1 h is allowed while filters are changed. Usually, the collection device should be changed at least once a day. The regulations allow 3 d to analyze each filter, though if there is a significant rise in turbidity, or if there are other reasons to suspect that oocyst counts may have risen, the filter should be changed and examined as soon as possible. The analysis has to be carried out in accredited laboratories using approved standard operating procedures. Laboratories have to participate in an external quality assurance scheme. As the results may be used to support a criminal prosecution, the guidance documents specify a forensic degree of chain of evidence, using tamper-proof containers and high levels of documentation. Results are reported to the DWI monthly, though abnormally high results must be reported immediately. These results are also in the public domain and are available to consumers should they wish to see them. Abnormally high results are to be reported to the relevant health authority, who will make an appropriate assessment of whether this poses a threat to public health and take the necessary steps to
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mitigate that threat. The U.K. Public Health Laboratory Service issued advice on the interpretation of these results when the regulations came into force (Hunter, 2000). Under these regulations, there were no outbreaks linked to drinking water in England and Wales until December 2005, when an outbreak linked to a reservoir in North Wales affected 231 people (Anon., 2005). Ironically, this supply was not part of the continuous monitoring scheme as it had not been classified as at risk.
B.
U.S. EPA Regulations
In the United States, public health protection from Cryptosporidium and many other contaminants is managed through a risk-based approach, where the frequency of occurrence of oocysts in a public water supplier’s source water is weighted against risk-of-infection probabilities in the consuming public to define treatment levels necessary to keep actual infection rates below a specified maximum level. Treatment may be accomplished either by removal (filtration), disinfection, or a combination of both. In 2006, the U.S. EPA finalized the Long Term 2 Enhanced Surface Water Treatment Rule (LT2) (USEPA, 2006a) to “improve the control of Cryptosporidium and other microbiological pathogens in drinking water systems with the highest risk levels.” The EPA estimates that full compliance with the LT2 will reduce the incidence of cryptosporidiosis—the gastrointestinal illness caused by ingestion of Cryptosporidium—by 89,000 to 1,459,000 cases per year, with an associated reduction of 20 to 314 premature deaths. The rule requires monitoring Cryptosporidium oocyst levels in the source water to establish the degree of log reduction (removal or disinfection) required to protect public health. Sources with higher oocyst concentrations will require more stringent treatment. The rule uses a treatment technique approach, assigning log credits to a wide range of processes based on their efficacy at removing or disinfecting Cryptosporidium. LT2 applies to all public water systems (systems) that use surface water or groundwater under the direct influence of surface water. This includes about 14,000 systems serving approximately 180 million people. LT2 requires utilities to monitor monthly for a period of 2 years to determine the mean Cryptosporidium concentration in the source water. The EPA has developed a list of approved laboratories that meet the criteria established for Cryptosporidium monitoring under the rule. Utilities must collect a minimum of 24 samples in the 2-year period on an approved schedule, but additional sampling is acceptable. The bin classification is based on total oocyst count determined by Method 1622. If a utility collects 48 samples, the mean concentration will be the 2-year mean; if 24 to 47 samples are collected, the mean used for bin determination will be the highest mean of samples in 12 consecutive months. The mean concentration will determine a category or bin into which the treatment plant falls, and each bin has an assigned level of treatment necessary to comply with the Rule. Table 11.2 shows the bins, the range of mean Cryptosporidium levels, and treatment required. The base level of treatment is conventional treatment. Treatment technologies are reviewed in Section V. Most systems are expected to be classified in the lowest bin and will face no additional requirements. Systems classified in higher bins must provide additional water treatment to further reduce Cryptosporidium levels by 90 to 99.7% (1.0 to 2.5-log), depending on the bin. Systems will select from different TABLE 11.2 Bin Classification for Treatment Under the Long Term 2 Enhanced Surface Water Treatment Rule Bin Number
Mean Cryptosporidium Concentration
Additional Treatment Requirements for Systems with Conventional Treatment
1 2 3
Oocysts < 0.075/L 0.075/L < oocysts < 1.0/L 1.0/L < oocysts < 3.0/L
4
Oocysts ≥ 3.0/L
No action 1-log (0.5 + 0.5 log or 1.0 log or greater from toolbox) 2.0 logs (with disinfection) with at least 1-log inactivation—e.g., UV, ozone, chlorine dioxide, membranes, bag filters, or river bank filtration 2.5 logs (with disinfection) with at least 1-log inactivation—e.g., UV, ozone, chlorine dioxide, membranes, bag filters, or river bank filtration
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treatment and management options in a “microbial toolbox” to meet their additional treatment requirements. All unfiltered water systems must provide at least 99 or 99.9% (2- or 3-log) inactivation of Cryptosporidium, depending on the results of their monitoring. In addition to the requirement for Cryptosporidium monitoring and perhaps additional treatment, LT2 requires that systems that store treated water in open reservoirs must either cover the reservoir or treat the reservoir discharge to inactivate 4-log virus, 3-log Giardia lamblia, and 2-log Cryptosporidium. Systems must also review their current level of microbial treatment before making a significant change in their disinfection practice. This review will assist systems in maintaining protection against microbial pathogens as they take steps to reduce the formation of disinfection byproducts under the Stage 2 Disinfection Byproducts Rule, a companion rule to LT2. Unlike the U.K. regulations, LT2 does not require monitoring of finished water nor does it set a maximum contaminant level that is met through monitoring (the maximum contaminant level goal is set at zero for Cryptosporidium). The goal of better public health protection is achieved through the use of established treatment practices, proper operational control, and easily measurable water quality parameters (turbidity) to ensure that treatment is working properly.
V.
Drinking Water Treatment
When the first outbreaks of cryptosporidiosis occurred, drinking water professionals were uncertain as to how to control Cryptosporidium in the water supply. The EPA had just promulgated the Surface Water Treatment Rule (USEPA, 1989) to control Giardia, and studies were undertaken to determine if oocysts were similar to cysts with respect to physical removal and inactivation. Cryptosporidium oocysts are much smaller than Giardia cysts, which could affect their ability to be filtered. Giardia cysts are much more resistant than bacteria and viruses to chlorine based disinfectants, but are susceptible at sustained contact time. Researchers scrambled to learn as much as possible about the ability of current treatment technologies to control Cryptosporidium as well as searching for alternative technologies (Schuler and Ghosh, 1990; Hall et al., 1994; Patania et al; 1995; Plummer et al., 1995; Edzwald and Kelly, 1998; Gregory et al., 1999; Medema et al., 2000; Wang et al., 2000). In promulgating LT2, the EPA conducted a thorough review of all of the technologies for treatment of Cryptosporidium in water, and as such the rule itself is the premier current reference on Cryptosporidium treatment; its literature reviews will be the reference foundation of much of the following discussion on treatment of Cryptosporidium in drinking water. Technologies to be covered are listed in Table 11.3, as are the range of log reduction credits available to water suppliers by employing these processes. Note that for some, such as chemical disinfectants, the actual inactivation credit may be dependent on water conditions such as pH or temperature. Chapter 1 discussed the efficacies of specific physical and chemical disinfectants in the context of the biology of Cryptosporidium and the hardiness of oocysts to environmental stresses, as well from the perspective of interrupting the host–parasite cycle for protection of public health. This chapter focuses on the practical application of many of those disinfectants and physical removal technologies (filtration) for control of the waterborne transmission route.
A. 1.
Filtration Technologies Conventional Filtration
Defined as filtration treatment consisting of coagulation, flocculation, sedimentation, and filtration, conventional filtration has been characterized in a number of studies as being capable of attaining at least 3 log or 99.9% removal of Cryptosporidium (McTigue et al., 1998; Patania et al., 1999; Huck et al., 2000; Emelko et al., 2000; Harrington et al., 2001; Nieminski and Bellamy, 2000; Dugan et al., 2001). That level was adopted by the EPA as the treatment credit granted under LT2 to water suppliers employing conventional filtration, or similar filtration substituting alternative clarification processes, including dissolved air filtration, for sedimentation. Nevertheless, outbreaks of cryptosporidiosis have
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TABLE 11.3 Cryptosporidium Treatment Processes with Available Log Credits Assigned by the EPA in the Long Term 2 Enhanced Surface Water Treatment Rule (LT2) Process/Technology
Log Credita
Conventional filtration Direct filtration Slow sand primary filtration Diatomaceous earth filtration Combined filter performance Individual filter performance Presedimentation basin with coagulation Two-stage lime softening Riverbank filtration Bag and cartridge filtration Membrane filtration Second-stage filtration Slow sand secondary stage filtration Chlorine dioxide Ozone UV
3.0 2.5 3.0 3.0 0.5 0.5 0.5 0.5 0.5–1.0b or >1.0c Up to 2.5c Per demonstrationc 0.5 2.5 Up to 3.0d Up to 3.0d Up to 4.0c,d
a
b c
d
Credit assumes performance of process within acceptable operational specifications. Credit dependent on process design. Attainment of credit requires preliminary process demonstration/validation. Credit dependent on design and water quality conditions (see inactivation tables, Tables 11.2–11.4)
Source: U.S. EPA Federal Register 40 CFR Parts 9, 141, and 142, National Primary Drinking Water Regulations, Long Term 2 Enhanced Surface Water Treatment Rule; Final Rule. 2006.
occurred in systems employing conventional treatment plants, including those in Milwaukee in 1993 (MacKenzie et al., 1994), and Waterloo, ON, in 1993 (Pett et al., 1993). Instances of high protozoa concentration in the source water or of poor filter performance may result in sufficiently high numbers of infectious oocysts in the finished water to cause an outbreak.
2.
Direct Filtration
Direct filtration is similar to conventional treatment in that a coagulant is used to form larger particles, but the settling or sedimentation step is not included. Water quality that is acceptable for direct filtration is generally of low and consistent turbidity, and coagulated water is applied directly to the filters. It is generally agreed that direct filtration will provide 2.5 log Cryptosporidium removal. In 1994, Las Vegas, NV, experienced a cryptosporidiosis outbreak in its direct filtration plant, which was a new facility and properly operated (Roefer et al., 1996). The EPA allows additional removal credits available to conventional and direct filtration systems by 0.5 log or greater by employing any of a number of supplemental treatment stages, including riverbank filtration, presedimentation and coagulation, and second-stage lime softening, or by operating within specified turbidity limits as measured at the combined filter effluent and/or at each individual filter.
3.
Slow Sand Filtration
This filtration process is credited with 3.0-log credit when used as a primary filtration, and 2.5-log in the secondary filtration role. Slow sand filtration uses a biological process to remove organic contaminants from the source water. A biological ecosystem, the schmutzdecke, forms on top of the sand filter and
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dissolves organic material as it passes through in the water. This process is nonpressurized and operates slowly as compared to other types of sand filtration. Pathogens including Cryptosporidium are removed in this treatment process (Schuler and Ghosh, 1991; Fogel et al., 1993; Timms et al., 1995). However, as noted with conventional treatment, the process may prove inadequate in the face of poor source water quality or deficient process performance. Nichols et al. (2001) have noted that slow sand filtration was implicated in several cryptosporidiosis outbreaks in the United Kingdom.
4.
Diatomaceous Earth Filtration
In this unique filtration process, a depth filter is developed after each backwash by applying granular media (diatomaceous particles) of specified size distribution to a backing screen while operating the filter in a recirculating mode. When sufficient media has been introduced and a filter cake has developed to the desired level, the filter is rinsed and placed in operation. Ongerth and Hutton (1997, 2001) demonstrate that at least 3-log removal can be expected, and with optimization even 6-log or higher may be achieved.
5.
Bag or Cartridge Filtration
The use of modular sieve-type filtration can be a very effective means of removing protozoa from drinking water. Bag filters are sheets of filter material secured around their perimeter in a loose manner such that the sheet extends as a pocket or bag in the direction of flow. Such a configuration offers greater surface loading area than would a tight filter sheet of the same diameter. Cartridge filters consist of surface or depth filter media wrapped or constructed around a hollow core, wherein the feed water contacts the outer perimeter and, with pressure, moves across the filter to the core. Media is often pleated to increase the effective filter surface area; in some cartridges, media of differential porosity is used, the outer depth being of higher pore size to trap larger particles, while the inner media is tighter, to capture finer material. In both filter types, inlet and outlet waters are separated by an O-ring, gasket, or mechanical seal to prevent bypassing of the filter media. Because the effectiveness of any bag or cartridge system is dependent on the characteristics of the filter media itself, and because these characteristics range widely, treatment credit is available only to the degree for which the filter effectiveness at removing Cryptosporidium or a surrogate has been demonstrated (including a 1-log safety factor) and only to a maximum of 2-log removal for a single bag or cartridge filter, or up to 2.5-log for two or more in series. Li et al. (1997) recognized that bag and cartridge performance can degrade with use, and current EPA regulations require that the pilot demonstration of any bag filter system be carried out across a range of periods of use, from new to end of run (terminal pressure drop).
6.
Membrane Filtration
In membrane filtration processes, water is forced by pressure (positive or vacuum) across a surface, excluding or rejecting particles greater than the effective pore size of the membrane material. Membrane filtration processes have been classified as reverse osmosis, nanofiltration, ultrafiltration, and microfiltration, with filter porosity correspondingly increasing from the molecular to the micrometer level. The distinction between these filters and the cartridge filters discussed earlier is that, to be recognized as membrane filters by the EPA, not only must representative specimens of filter models undergo demonstration testing of Cryptosporidium removal efficacy, but individual filters must also be capable of being integrity tested over the course of their use. Nondestructive tests, such as diffusive-flow and pressurehold tests (Meltzer, 1993), allow routine and regular (even daily) evaluation of individual filter units. The value imparted by direct integrity testing is such that the ceiling on removal credit is increased from the 2-log (99%) credited to cartridge filters to as high as 6-log (99.9999%) for membrane filtration.
B.
Disinfection
Disinfection is a form of treatment in which the biological contaminant is not necessarily removed from the water but, rather, is inactivated and rendered unable to cause infection. Although chlorine is the
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most commonly used disinfectant, Cryptosporidium oocysts proved to be resistant to chlorination at concentrations suitable for producing potable water. Ozone became the focus of much of the research in the early 1990s, but it was almost by accident that the most effective treatment—ultraviolet light—was discovered. Low doses of UV light are extremely effective in oocyst inactivation, and the use of UV treatment has become a large focus of the control of Cryptosporidium in water supplies.
1.
Ozone
The application of chemicals for a given level of disinfection is measured not in units of concentration alone, but as concentration over time. Chemicals may, in fact, be added to the water in measured concentrations, but the objective is to achieve a concentration over a sufficient period of contact, or concentration × time, or CT, which is expressed as mg/L × min. Ozone, similar to other oxidants, must be dosed in excess of the theoretical concentration necessary to achieve a certain CT, due to the presence of dissolved and suspended materials, especially organic species, which exhibit an ozone demand. Organic materials diminish the initial ozone residual to some extent, steadily interact with the oxidant at a water quality dependent decay rate and, eventually (generally in minutes), totally deplete the residual ozone. Thus, the overall CT for ozone is not the product of the applied concentration and contact time, but the integrated value for the concentration following initial demand through the end of the recognized contact zone or to the point where no residual remains. A given chemical disinfectant CT, regardless of whether it is applied with a high dose for a short period of time, or a low dose for a proportionally longer period of time, will effect the same disinfection as long as it is applied in the same physical/chemical environment. For ozone, the environmental factor most affecting the efficacy of a given CT is temperature, with reaction rates—and so inactivation kinetics—decreasing steadily with temperature. Ozone (O3) is an unstable oxidant, and so is generated on site in the gas phase by oxidation of pure oxygen or oxygen in air, then fed immediately into the water stream through a diffuser to maximize the gas–water interface. A common approach to dissolving the gas in water is to feed the gas into a contactor at the bottom of a downward flowing column of water, so that the ozone bubbles travel against the flow, dissolving as they travel. Although used effectively to control taste and odor, ozone can also be used for oocyst inactivation (Finch et al., 1993, 1997; Finch and Belosevic, 1999), but it is not as effective as initially thought. Early studies by Finch and Belosevic in demand-free water or buffer indicated greater inactivation than when natural matrices were used (Oppenheimer et al., 2000). After assessing these and a number of other studies (Rennecker et al., 1999; Li et al., 2001; Owens et al., 2000; Oppenheimer et al., 2000), the USEPA determined the relationship between ozone CT and the log inactivation of Cryptosporidium to be defined by the equation, Ozone Log (inactivation) credit = (0.0397) × (1.09757)Temp × CT where Temp is the water temperature in degrees centigrade (USEPA, 2006b). From this equation is derived the ozone CT table (see Table 11.4) displaying the log credits granted for ozonation across a range of CTs and across a range of temperatures. As temperatures approach freezing, the CT rises dramatically, to the point where six times the ozone CT is specified in the CT table for a given log inactivation at less than 0.5˚C, compared to that needed at 20˚C. Caution is recommended at lower temperatures; however, as several studies of ozone disinfection of Cryptosporidium in cold water indicate, the log inactivations shown in the CT tables in fact may not be achievable in certain waters at temperatures of 3˚C or less (Li et al., 1999; Oppenheimer et al., 2000).
2.
Chlorine Dioxide
Like ozone, chlorine dioxide (ClO2) is an unstable oxidant generated on site, though not as a gas but as a dilute liquid. Chlorine dioxide is generated by reacting sodium chlorite with a strong oxidant, which may be gaseous or aqueous chlorine, mineral acid with or without chlorine, or acid with a hypochlorite solution (Gates, 1998). Also like ozone, the efficacy of chlorine dioxide for the disinfection of Cryptosporidium is strongly dependent on temperature. Based on the studies of Li et al. (2001), Owens et al.
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TABLE 11.4 Ozone CT Values (mg/L × min) Necessary for Cryptosporidium Inactivation Credit Water Temperature (°C) 5 7 10
Log Credit
≤ 0.5
1
2
3
0.25 0.5 1.0 1.5 2.0 2.5 3.0
6.0 12 24 36 48 60 72
5.8 12 23 35 46 58 69
5.2 10 21 31 42 52 63
4.8 9.5 19 29 38 48 57
4.0 7.9 16 24 32 40 47
3.3 6.5 13 20 26 33 39
2.5 4.9 9.9 15 20 25 30
15
20
25
30
1.6 3.1 6.2 9.3 12 16 19
1.0 2.0 3.9 5.9 7.8 9.8 12
0.6 1.2 2.5 3.7 4.9 6.2 7.4
0.39 0.78 1.6 2.4 3.1 3.9 4.7
Source: USEPA. Ultraviolet Disinfection Guidance Manual Office of Water EPA 815-R-06-007. Washington, D.C. 2006.
TABLE 11.5 Chlorine Dioxide CT Values (mg/L × min) Necessary for Cryptosporidium Inactivation Credit Water Temperature (°C) 5 7 10
Log Credit
≤ 0.5
1
2
3
0.25 0.5 1.0 1.5 2.0 2.5 3.0
159 319 637 956 1275 1594 1912
153 305 610 915 1220 1525 1830
140 279 558 838 1117 1396 1675
128 256 511 767 1023 1278 1534
107 214 429 643 858 1072 1286
90 180 360 539 719 899 1079
69 138 277 415 553 691 830
15
20
25
30
45 89 179 268 357 447 536
29 58 116 174 232 289 347
19 38 75 113 150 188 226
12 24 49 73 98 122 147
Source: USEPA. Ultraviolet Disinfection Guidance Manual Office of Water EPA 815-R-06-007. Washington, D.C. 2006.
(1999), and Ruffell et al. (2000), the USEPA (2006b) adopted the log (inactivation) credit formula for chlorine dioxide disinfection of Cryptosporidium to be ClO2 Log (inactivation) credit = (0.001506 × (1.09116)Temp × CT The resultant CT table derived from this equation is shown in Table 11.5. Thus, chlorine dioxide requires a much higher CT than does ozone for a given log inactivation of Cryptosporidium, but the impact of temperature on both oxidants is quite similar.
3.
Ultraviolet Light (UV)
UV is unlike any of the chemical disinfectants, in that it is a physical process whereby photons of a germicidal wavelength (200 to 300 nm, and particularly about 260 nm) are absorbed by molecules of an irradiated organism, causing damage to the organism. Specific to disinfection is the absorption of UV photons by base pairs of the DNA, producing, as a photo by-product, a dimerization of the damaged pairs. A consequence of this damage is that subsequent cell replication is interrupted, and so host infection is prevented. UV is typically generated as an emission from electrically excited mercury vapor or xenon molecules, which, depending on the nature of the lamps, may be nearly monochromatic, as is the output of low mercury pressure lamps, at 254 nm, or polychromatic, as are the outputs of medium-pressure mercury lamps and xenon lamps. In drinking water applications, UV reactors containing 1 to over 100 lamps are housed in a closed reactor through which the water to be treated flows. In units of more than rudimentary complexity, a UV sensor responds to the irradiation from the lamp, with sensor readings increasing in proportion to lamp output and inversely to the UV absorbance of the water.
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The particular history of UV disinfection of Cryptosporidium obligingly follows the pattern noted in understanding Cryptosporidium treatment by any means: studies were generally lacking until a public health threat was perceived. Details of this history are of interest, however, and will be examined here to a degree beyond the level given to other technologies. This scrutiny is justified as well by the degree of effectiveness of UV technology. Long before any Cryptosporidium outbreaks occurred, UV had been known to be an effective agent against bacteria. As early 1966, UV was accepted by the U.S. Department of Health, Education, and Welfare as a means of disinfecting bacteria in drinking water, applied in a minimum dose of 16 mJ/cm2. Research into the treatability of viruses had shown these pathogens to be susceptible to UV irradiation, which came to be employed broadly to that end in wastewater applications. An obstacle to more numerous applications of UV to drinking water, however, was that it was not chlorine. That chemical disinfectant was widely used in the 20th century, had few known drawbacks, was relatively easy to apply, and left a residual. This latter characteristic offered two significant advantages over UV: the residual achieved downstream disinfection from the point of application, and straightforward measurement of disinfection levels. In addition, the protozoan parasite of concern in the mid to late 20th century, Giardia, was susceptible to chlorination, which held its primary disinfectant position with little competition. After a number of cryptosporidiosis events in the United States, Cryptosporidium disinfection research began in earnest. The widely applied chlorine was found to be ineffective against Cryptosporidium oocysts (Korich et al., 1990). Just before the Milwaukee outbreak occurred, a study by Ransome et al. (1993) evaluated disinfection of Cryptosporidium treated by the chemical agents chlorine, ozone, and chlorine dioxide, among others, and by UV. Assaying the treated oocysts using excystation, the study results corroborated those of Korich et al. (1990), by indicating little effectiveness with chlorine. The results also demonstrated that a 1-log reduction in viability was achieved by an ozone CT of 1.8 mg min/L, a chlorine dioxide CT of 75 mg min/L, and a UV dose of 80 mJ/cm2, with 120 mJ/cm2 achieving 2-log inactivation. These latter values were 5 to 10 times the UV dose understood to be highly effective against bacteria, and over twice that generally applied for control of viruses. The authors dismissed UV for Cryptosporidium disinfection, inasmuch as one would need to apply a dose beyond that called for by a number of UV guidelines then in effect. Citing the poor performance of chlorine in the face of cryptosporidiosis outbreaks, and recognizing there might actually be a role for UV against the protozoa, Campbell et al. (1995) challenged a low-pressure mercury lamp UV system with Cryptosporidium oocysts, applying over 8000 mJ/cm2 in the unit being evaluated. Again, using excystation to measure effect, the researchers demonstrated over 2-log inactivation (the upper limit was not determined), and stressed that the actual log inactivation that may have been achieved was limited not by the process but by the assay methodology. The report ends with a recommendation for an evaluation of UV using a more sensitive assay. Clancy et al. (1998) took that next step, testing the same UV system and assessing its effect using animal infectivity, and demonstrated that the UV system applying 4000 to 8000 mJ/cm2 had achieved over 4-log inactivation (upper limit also not determined), flowing at 100 to 400 gal/min (gpm). In the same time period, a comparison of in vivo animal infectivity methods against in vitro (excystation and vital dye stains) viability methods revealed that UV, in doses as low as 14 mJ/cm2, in this case delivered from a pulsed xenon lamp, achieved greater than 3-log inactivation as measured by animal infectivity (Clancy et al., 2000c; LaFrenz, 1997), whereas viability assays indicated the same UV had no appreciable effect. Building on these findings, a study of medium pressure UV’s effect on Cryptosporidium (Bukhari et al., 1999) and a follow-up study using low pressure UV (Clancy et al., 2000d) revealed that UV could achieve 4-log inactivation or greater at doses as low as 10 mJ/cm2. To this point, much of the infectivitybased research on low dose UV inactivation of Cryptosporidium had been performed by a single team, but by 2001, others had corroborated these findings (Craik et al., 2001; Shin et al, 2001). A shortcoming of this growing body of knowledge was that all the data were generated using a single isolate, the Iowa strain. To remedy this, Clancy et al. (2002) demonstrated UV was similarly effective against all five strains of C. parvum tested (Glasgow, Iowa, Maine, Moredun, Texas A&M). Other research determined that Cryptosporidium oocysts do not have the capacity for repair following UV irradiation (Shin et al., 2001, Rochelle et al., 2004). A summary of UV disinfection is presented in Table 11.6.
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TABLE 11.6 Research Studies on the Effect of UV on Cryptosporidium Water Matrix
UV Type/System Scale
UV Dose (mJ/cm2)
Log Inactivation In vitro 1 2 >2
Log Inactivation In vivo
Study
Organism
Ransome et al., 1993 Campbell et al., 1995 Dunn et al., 1995 Finch et al., 1997 Clancy et al., 1998 Bukhari et al., 1998 Bukhari et al., 1999
C. parvum
NA
LP/bench
C. parvum
Clean
LP/pilot
80 120 8,700
C. parvum
NA
PUV/bench
1000
>6
C. parvum Iowa C. parvum Iowa C. parvum Iowa C. parvum Iowa
DI
LP/bench
41,000
< 0.6
Drinking water
LP/1 gpm LP/400 gpm PUV/bench
180 8,700 40
<1 >2 0.3
Drinking water
MP/bench 200 gpm
< 0.1
Finch and Belosevic, 1999
C. parvum Iowa G. muris
Drinking water
MP/bench
19 60 159 8
Drinking water
MP/bench
0.1 0.1
Clancy et al., 2000a Clancy et al., 2000c Klevens, 2001
DI DI
LP/bench MP/bench PUV/bench
Linden et al., 2001 Mofidi et al., 2001
C. parvum Iowa C. parvum Iowa C. parvum Iowa C. parvum Iowa C. parvum Iowa
21 83 9 9 40 14 50 50 3
Mackey et al., 2002 Clancy et al., 2002
DI
Dilute PBS
LP MP LP/bench
Filtered surface water
PUV and MP/bench
C. parvum Iowa C. parvum Iowa
Drinking water
LPHO/full scale
TAMU
DI
Maine Moredun
Drinking water
LP/bench
>4 > 2.8 3.9 >4 3.2 2.8 2.8 3.5 3.6 > 2.5 > 2.5 > 5.7 > 5.7 > 3*
3 6 9 45
1.0* 1.9* 2.7* > 4.7
10 4 2 40 10 20 5 10
>4 4 3 > 5.4 5 6.0 4.3 > 5.7
* Cell culture method. Note: In vivo refers to neonatal mouse infectivity assay unless otherwise noted. In vitro assays are excystation and/or vital dye staining. LP = low pressure UV; MP = medium pressure UV; PUV = pulsed UV; LPHO = low pressure high output UV.
With this information, the EPA recognized UV as a means by which water utilities with surface water sources could attain improved Cryptosporidium reduction while minimizing the disinfection byproducts resulting from the use of chlorine and ozone. Several researchers had demonstrated a similar sensitivity of Giardia to UV as well (USEPA Draft Ultraviolet Disinfection Guidance Manual [UVDGM], 2003). The end result of these studies was that the EPA included UV as a prominent tool for drinking water
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TABLE 11.7 UV Dose Necessary for Cryptosporidium Inactivation Credit Log Credit 0.5 1.0 1.5 2.0 2.5 3.0 3.5 4.0
Cryptosporidium UV Dose (mJ/cm2) 1.6 2.5 3.9 5.8 8.5 12 15 22
Giardia UV Dose (mJ/cm2) 1.5 2.1 3.0 5.2 7.7 11 15 22
Source: USEPA. Ultraviolet Disinfection Guidance Manual Office of Water EPA 815-R-06-007. Washington, D.C. 2006.
utilities to utilize in meeting their disinfection goals while minimizing disinfection byproduct formation. UV dose tables, shown in Table 11.7, were prepared for up to 4-log inactivation of these protozoa (USEPA, 2006b). Because UV irradiation does not provide a residual for dose delivery verification, the EPA has required utilities using UV to install systems whose performance and monitoring systems have been validated. Extensive information on UV technology, including reactor validation is provided by the EPA in the Final UVDGM (USEPA, 2006b). With a validated reactor, a utility will have assurance that a system can achieve the needed level of disinfection for the log credit being sought, and that the monitoring system can be relied upon to indicate when the system is operating within specification and when it is not.
VI. Conclusions About two decades have passed since Cryptosporidium was recognized as a human waterborne pathogen, capable of causing serious morbidity and even mortality in individuals consuming tap water. In that short period, a tremendous amount of energy was directed toward understanding the occurrence of Cryptosporidium in water, and how to control it through water treatment. Dozens of studies have determined that Cryptosporidium oocysts can be found in the environment worldwide, in source and treated drinking water. Although methods such as Method 1622 are significant improvements over previous testing methods, issues remain with specificity and sensitivity. A host of molecular techniques are available and will become part of the standardized testing protocols over the next decade (see Chapter 5 for the usage of genotyping tools in assessing the source and human infection potential of Cryptosporidium oocysts in water). After its role in drinking water contamination was recognized, the inability of conventional water treatment to control the parasite in water supplies was a major challenge to water treatment professionals. Infectious oocysts that passed through the filtration process were unaffected by chlorine and chlorinebased disinfectants, and outbreaks occurred even in plants meeting all water quality and operational standards. The use of UV technologies for Cryptosporidium treatment in drinking water is a major breakthrough for the water industry. UV was rapidly adopted by the EPA for treatment of surface waters used as drinking water sources, and is gaining popularity worldwide for Cryptosporidium control. Cryptosporidium testing has a role and utilities will continue to test to better understand their risk levels, but prevention of waterborne disease and protection of public health is the result of proper drinking water treatment, storage, and transmission, not pathogen testing.
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12 Waterborne: Recreational Water*
Michael J. Beach
CONTENTS I. II.
Introduction .................................................................................................................................. 335 Factors Contributing to the Transmission of Cryptosporidium through Recreational Water Use................................................................................................................ 336 III. Case Surveillance and Epidemiologic Studies of Associated Risk Factors for Cryptosporidiosis.................................................................................................................... 339 IV. Outbreaks of Cryptosporidiosis Associated with Recreational Water Use ................................. 341 A. Outbreaks in the United States: General Overview ....................................................... 341 B. Outbreaks in Countries Other than the United States: General Overview .................... 350 C. Recreational Water Types and Outbreak-Associated Deficiencies................................. 350 1. Marine Water.................................................................................................... 350 2. Freshwater Lakes, Rivers, and Hot Springs .................................................... 351 3. Disinfected Water............................................................................................. 352 V. Recreational Water Types and Prevention ................................................................................... 357 A. Marine and Fresh Water.................................................................................................. 357 B. Disinfected Water ............................................................................................................ 358 VI. Perspective.................................................................................................................................... 360 Acknowledgments.................................................................................................................................. 361 References........................................................................................................................... 361
I.
Introduction
Recreational water—which includes natural water found in oceans, lakes, rivers, and hot springs, as well as artificial, disinfected venues such as swimming pools, water parks, hot tubs, and interactive fountains—has been well documented through outbreaks and epidemiologic studies as a transmission vehicle for pathogens (Stevenson, 1953; Cabelli et al., 1979; Dufour, 1984; Seyfried et al., 1985; Cheung et al., 1990; Ferley et al., 1989; Fleisher et al., 1998; Pruss, 1998; Haile et al., 1999; Henrickson et al., 2001; Craun et al., 2005; Dziuban et al., 2006; Wade et al., 2006; Weidenmann et al., 2006). As a result, recreational water use has been associated with a spectrum of symptoms including skin (Gustafson et al., 1983; Tate et al., 2003; Dziuban et al., 2006), ear (Havelaar et al., 1983; van Asperen et al., 1995), eye (D’Angelo et al., 1979; Papapetropoulou and Vanatarakis, 1998), respiratory (Jernigan et al., 1996; Fields et al., 2001), neurologic (Visvesvara and Stehr-Green, 1990; CDC, 2004) and, most commonly, acute gastrointestinal illness (AGI; Pruss, 1998; Dziuban et al. 2006, Smith et al., 2006; Wade et al., 2006). Since its identification as a human pathogen in 1976 (Meisel et al., 1976; Nime et al., 1976), Cryptosporidium has emerged as the major cause of outbreaks of AGI associated with use of recreational water, particularly those outbreaks associated with artificial, disinfected venues. This chapter summarizes * The findings and conclusions in this report are those of the author and do not necessarily represent the views of the Centers for Disease Control and Prevention.
335
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Cryptosporidium and Cryptosporidiosis, Second Edition TABLE 12.1 Factors Facilitating Transmission of Cryptosporidium in Recreational Water Settings Parasite 1. 2. 3. 4. 5.
Transmission route simple (i.e., fecal oral). Oocysts immediately infectious. Low infectious dose. High titer of oocysts in stool. Excretion in stool prolonged and continues after cessation of diarrhea.
Environment (oocyst) 1. 2. 3. 4.
5. 6.
Persistent in the environment (fresh and marine water, low temperatures). Viable oocysts found in treated wastewater. Transmission can be zoonotic. Contamination of recreational water by feces is ubiquitous. a. Animals b. Rainfall run-off events c. Wastewater outflows d. Sewer overflows e. Fellow swimmers i. Direct fecal accidents ii. Residual feces on swimmers bodies, poor hygiene Small size challenges filtration systems. Resistant to halogen disinfection.
Population 1. 2. 3. 4. 5. 6.
Diarrhea is common in population. Cryptosporidiosis is an endemic disease in population. Antibody appears to confer protection against severe illness rather than infection. a. Reinfection can occur Recreational water exposure is high. Swimming pool operation is not optimal. Lack of awareness in the general public about pathogens spread via recreational water. a. Water swallowing while swimming is common b. Hygiene improvements needed c. Swimmers continue swimming when ill with diarrhea
(1) the factors facilitating transmission of Cryptosporidium via recreational water use, (2) data from case surveillance and epidemiologic studies, (3) data from outbreak reports, and (4) changes in operation and use of recreational water that are needed to reduce the risk of future transmission.
II.
Factors Contributing to the Transmission of Cryptosporidium through Recreational Water Use
There are multiple parasite, environmental, and populational factors that make Cryptosporidium ideally suited for transmission through recreational water, which may also be examined in other chapters of this book (Table 12.1). The parasite’s simple fecal-oral transmission route is optimal when combined with a water-based vehicle in which large numbers of people of all ages commonly immerse themselves and swallow the water, as with recreational water. Oocysts are immediately infectious, and the parasites’ infectious dose, 10 to 30 oocysts, is low (DuPont et al., 1995, Okhuysen et al., 1999). Oocysts are excreted in large quantities in animals (from 105 to 107 oocysts/gram of stool; Current, 1985; Uga et al., 2000) and humans (≤ 109 oocysts in stool per day; (Jokipii and Jokipii, 1986; Goodgame et al., 1993;
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Chappell et al., 1996). Excretion of oocysts for up to 50 days after cessation of diarrhea (Jokipii and Jokipii, 1986; Stehr–Green et al., 1987; Chappell et al., 1996) also increases the risk of transmission when swimmers return to the water following resolution of their diarrhea. Oocysts are environmentally persistent and can remain infectious for months to a year in either freshwater (Robertson et al., 1992; Pokorny et al., 2002) or seawater (Robertson et al., 1992; Fayer et al., 1998a; Tamburrini and Pozio, 1999), and longer at colder temperatures (Fayer and Nerad, 1996; Fayer et al., 1998b; Tamburrini and Pozio, 1999; Pokorny et al., 2002). Surface water becomes contaminated by zoonotic sources of oocysts that are readily mobilized by precipitation events in the surrounding watershed. In addition, because oocysts can be found in treated wastewater (Xiao et al., 2001; Ward et al., 2002; Montemayor et al., 2005), regular releases of treated or untreated wastewater through treatments plants, combined sewer overflows, and sanitary sewer overflows routinely contaminate recreational water sources. As a result, Cryptosporidium is common in the environment and surveys of surface water sources from numerous countries routinely detect Cryptosporidium (Rose, 1988; LeChevallier et al., 1991; LeChevallier and Norton, 1995; Xiao et al., 2001; Ward et al., 2002; Montemayor et al., 2005; Coupe et al., 2006; Karanis et al., 2006). Direct human contamination of recreational water can also occur. High levels of oocysts in stool make it possible for a single ill person’s bowel movement (108 to 109 oocysts can be released; Jokipii and Jokipii, 1986; Chappell et al., 1996) to significantly contaminate beaches and even multimillion gallon, artificial venues such as water parks. In addition, poor hygiene practices mean that most swimmers retain some level of feces on their perianal surface that can be rinsed into recreational water while swimming (Gerba, 2000). Although relatively small amounts of fecal contamination per person have been documented (0.14 grams per person, range 0.01 to 10 grams; Gerba, 2000), large, heavily used locations (e.g., 15,000 visitors per day) may receive daily fecal contamination loads on the order of 2.1 kg (4.6 lb) each day. Some of those individuals are likely to be excreting pathogens. One study in the United States found that 13/293 (4.4%) formed fecal accidents were positive for Giardia, although no Cryptosporidium was found (CDC, 2001b). Cryptosporidium oocysts and Giardia cysts have been detected in swimming pools in the absence of outbreaks of cryptosporidiosis or giardiasis (Fournier et al., 2002; Bonadonna et al., 2004; Greinert et al., 2004; Schets et al., 2004; Oliveri et al., 2006). A 1-year Dutch survey of filter backwash samples from five swimming pools found that 18/153 (11.8%) samples were positive for either Cryptosporidium (4.6%), Giardia (5.9%), or both (1.3%), making it likely that transmission via swimming is relatively common (Schets et al., 2004). For the artificial, disinfected venues, the parasites’ small size is a challenge for filtration systems (Rose et al., 1997). Most notable, however, is Cryptosporidium’s extreme resistance to halogen disinfection (Korich et al., 1990; Carpenter et al., 1999), which allows the parasite to bypass the major barrier to infectious disease transmission that has been used for the past 80 years in the recreational water industry. Typical state or local swimming pool codes in the United States require 1 ppm free residual chlorine in public pools (National Recreation and Park Association, 1999). This concentration enables Cryptosporidium to survive for over 6 days using a CT inactivation number of 9600 (Korich et al., 1990). Diarrhea is a common illness globally. In developed countries, studies estimate that 0.1 to 3.5 diarrheal episodes occur per person per year (Roy et al., 2006). Surveys in the United States indicate that 7.2 to 9.3% of the general public has had diarrhea in the previous month (Jones et al., 2007). Fecal incontinence is relatively common, with 2.2% of persons interviewed in a community-based Wisconsin survey having fecal incontinence; 70% of those persons being under the age of 65 (Nelson et al., 1995). A meta-analysis of anal incontinence studies demonstrated that the prevalence among people aged 15 to 60 was 0.8% in men and 1.6% in women (Pretlove et al., 2006). In addition, many recreational water facilities cater to families, which include a key incontinent population, diaper-aged children, who can easily contaminate recreational water. In developed countries, Cryptosporidium has been found in 1 to 2% of diarrheic stools; 3 to 5% in children (Cordell and Addiss, 1994; Casemore et al., 1997). High percentages of oocyst-positive children have been found to be asymptomatic in some studies (Cordell and Addiss, 1994). Serologic tests for Cryptosporidium demonstrate that antibody to the parasite is common, with almost 70% of individuals reaching positivity by age 70 (Frost et al., 2004; Casemore, 2006). Reinfection can occur, although symptoms are usually less severe (Casemore, 2006). These data support the endemic nature of cryptosporidiosis and that the population is routinely exposed to the parasite. In particular,
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young children have a higher incidence of symptomatic infection that increases their chance of contaminating recreational water (Hlavsa et al., 2005). Exposure to recreational water is massive and common. An estimated 7.4 million swimming pools are in residential or public use in the United States, and approximately 670,000 pools exist in Canada (Pool and Spa Marketing, 2003, 2006). In the United States, over 360 million annual visits to recreational water venues such as swimming pools, spas, lakes, rivers, and the ocean occur, making swimming the second most popular activity in the United States and the most popular activity for children (United States Bureau of the Census, 1995). Growth of water parks has been substantial in the past 20 years, so that currently there are over 1000 water parks in North America and 600 more, globally. Attendance at North American water parks in 2004 was approximately 73 million visits with 3 to 5% growth over the past 5 years (World Waterpark Association, 2006). Public swimming pool operation is usually subject to some level of government regulation, which usually results in periodic inspections to ascertain that aquatics facilities are maintaining regulatory compliance with existing pool codes. Surveys of pool inspection data in the United States (CDC, 2003) or pool water sampling in England and Wales (PHLS, 2003a; PHLS, 2003b) routinely demonstrate that a significant percentage of pools are found to be out of compliance with regulatory standards. In a survey of 22,131 pool inspection reports, 54.1% of inspections found violations and 8.3% of inspections resulted in the pool being immediately closed because of public health concern (CDC, 2003). From 4.5 to 18.4% of inspections documented violations of disinfectant level standards, with the highest percentage of violations being found with shallow children’s pools (CDC, 2003). Such shallow pools may be more difficult to maintain because of the shallow water and ease with which disinfectant is depleted because of depth, aeration, sunlight, and organic loads from young children. Unfortunately, these pools are frequented by a population that has a higher prevalence of Cryptosporidium and may experience more severe disease (Hlavsa et al., 2005); the pools with the highest potential risk for contamination have the poorest water quality. These data illustrate the challenges involved in enforcing pool regulations to ensure that trained staff are operating pools according to mandated water quality guidelines. This issue is compounded by the halogen resistance of Cryptosporidium so that even a well-maintained pool can be subject to contamination by, and transmission of, Cryptosporidium. The swimming public is generally unaware of recreational water illness issues that are needed to assist in reducing the level of pathogen transmission. In 1998 and 1999, the U.S. Centers for Disease Control and Prevention (CDC) conducted focus groups consisting of parents of young children who used recreational water parks (Macro International, Inc., 2000). Several conclusions were drawn from these focus groups regarding parents’ knowledge and attitudes about swimming that shed light on existing problems with treated water venues. These parents did not consider swimming as communal bathing or think about the shared nature of the water used for swimming. They believed that waterborne disease was something one finds only in other countries, and they were not aware of the potential for pool water to spread illness. One reason for not perceiving recreational water as a problem was that they believed chlorine disinfection of pools killed all pathogens immediately so that pool water was essentially “sterile.” Such beliefs likely underlie the common practice of swallowing recreational water (Dufour et al., 2006), poor hygiene (Gerba, 2000), and continued swimming during diarrheal illness that leads to transmission of recreational water illnesses. Although swimming is an active and healthy way to spend leisure time, the enormous popularity of the activity has created public health challenges for recreational water use. The combination of (1) Cryptosporidium’s biology and environmental hardiness, (2) the high level of exposure of all ages, particularly young children, (3) the endemic nature of the disease, (4) the poor hygiene of the general public, and (5) the low level of public awareness about recreational water illnesses has served as the foundation for Cryptosporidium’s becoming a major recreational-water-associated pathogen in developed countries (Table 12.1).
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III. Case Surveillance and Epidemiologic Studies of Associated Risk Factors for Cryptosporidiosis Case surveillance data are consistent with swimming as a significant risk factor for transmission. In the United States, cryptosporidiosis became a nationally notifiable disease in 1994. From 1995 to 2002, CDC received 24,312 cryptosporidiosis case reports with 3,787 in 2001 and 3,016 in 2002 (Dietz and Roberts, 2000; Hlavsa et al., 2005). The data reported likely underestimate the cryptosporidiosis burden in the United States. Enteric infections are highly underreported because (1) not all infected persons are symptomatic, (2) those who are symptomatic do not always seek medical care, (3) health-care providers do not always include diagnostics in their workup of diarrheal diseases because they might treat patients without testing stool for the pathogen, and (4) case reports are not always completed for positive laboratory results or forwarded to public health officials. CDC estimates that up to 300,000 cases per year occur based on documented underreporting of enteric diseases (Chalker and Blaser, 1988; Mead et al., 1999; Hlavsa et al, 2005; Jones et al., 2007). Even these estimates may be low considering the high antibody prevalence in the population (Frost et al., 2004; Casemore, 2006). Reporting of cryptosporidiosis in the United States demonstrates that cryptosporidiosis affects persons in all age groups (Figure 12.1; Hlavsa et al., 2005). However, the number of reported cases was highest among children 1 to 9 years of age and adults 30 to 39 years of age; two key groups using recreational water venues. In the United States, a marked seasonality in the onset of illness occurs in early summer through early fall with a fourfold increase in transmission (Figure 12.2). The seasonality is most marked in children 1 to 9 years old where five- to sevenfold increases occur (Figure 12.3). This increase coincides with increased outdoor activities and swimming during the summer recreational water season. This single summer seasonal peak has also been noted in Canada (Majowicz et al., 2001), Australia (Black and McAnulty, 2006), and New Zealand (Learmonth et al., 2003). Surveillance data from England and Wales has demonstrated a bimodal distribution with peaks in the spring and summer; the spring peak declined following the foot-and-mouth epidemic and improvements in public water supplies in the United Kingdom (Sopwith et al., 2005).
5000 4500 Number of cases
4000 3500 3000 2500 2000 1500 1000 500 0 under 1
1-4
5-9
10-14 15-19 20-24 25-29 30-34 35-39 40-44 45-49 50-54 55-59 60-64 65+*
*Case reports decreased with increased age.
Age group (yrs) FIGURE 12.1 Number of cryptosporidiosis case reports (n = 24,312) by age group—United States, 1995–2002.
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Number of cases
2000 1500 1000 500 0
J
F
M
A
M
J
J
A
S
O
N
D
Month FIGURE 12.2 Number of cryptosporidiosis case reports (n = 24,312) by date of illness onset—United States, 1995–2002.
600
1-4 years 5-9 years 30-34 years
Number of cases
500 400 300 200 100 0
J
F
M
A
M
J
J
A
S
O
N
D
Month FIGURE 12.3 Number of cryptosporidiosis case reports (n = 24,312) by selected age group* and date of illness onset—United States, 1995–2002. The 1 to 4 and 5 to 9 year age groups are presented because they have the highest numbers of cryptosporidiosis case reports and have the greatest seasonality. The 30 to 34 year age group was chosen to illustrate the less pronounced seasonality of the other age groups.
Epidemiologic case-control studies of sporadic cryptosporidiosis have also determined that recreational water is a risk factor in addition to other routes of transmission. In the United States, univariate analysis found freshwater swimming, marine swimming, and nonpublic pool swimming were significant risk factors; swimming in a home pool was protective (Roy et al., 2004). However, only freshwater swimming was implicated in the final multivariate model (Roy et al., 2004). In Australia, swimming in a public pool was implicated in a multivariate model (Robertson et al., 2002). In the United Kingdom, use of a toddler pool and the number of times one swam in a swimming pool (not use of a swimming pool) or toddler pool were significant in univariate analyses (Hunter et al., 2004). However, following multivariate analysis, swimming exposures were not significant. This case-control study was supported with genotyping data to differentiate between C. parvum and C. hominis. A multivariate model including only C. hominis did not find swimming exposures significant although the number of times one swam in a toddler pool had a p value of 0.077 (Hunter et al., 2004). An increase in the serologic response to Cryptosporidium antigens was used as a method to investigate risk factors associated with Cryptosporidium infection
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(Frost et al., 2000; Egorov et al., 2004). A serologic survey of college students in the United States before and after the introduction of a new community water treatment plant demonstrated that “swimming in a lake, stream, or pool” predicted an 18% increase in antibody response (Frost et al., 2000). In a similar study of Russian swimmers, males who swam in indoor pools had higher baseline antibody levels (Egorov et al., 2004). The observed increased tendency for males to put their head under the water was suggested as an explanation for the difference with female swimmers.
IV.
Outbreaks of Cryptosporidiosis Associated with Recreational Water Use
The first recreational-water-associated outbreaks of cryptosporidiosis were documented in 1988 and both outbreaks, one each in the United States and the United Kingdom, involved swimming pools (CDC, 1990; Sorvillo et al., 1992; Joce et al., 1991). Since 1988, cryptosporidiosis outbreaks associated with recreational water use have continued to be documented in the United States (n = 68; CDC, 1990; Sorvillo et al., 1992; Herwaldt et al., 1991; Moore et al., 1993; Kramer et al., 1996; Levy et al., 1998; Barwick et al., 2000; Lee et al., 2002; Yoder et al., 2004; Dziuban et al., 2006), Australia (n = 6; Lemmon et al., 1996; Beers et al., 1998; Hellard et al., 2000; Stafford et al., 2000; Puech et al., 2001; Black and McAnulty, 2006), Canada (n = 3; Bell et al., 1993; Macey et al., 2002; Louie et al., 2004), Japan (n = 2; Yokoi et al., 2005; Ichinohe et al., 2005), New Zealand (n = 1; Baker et al., 1998), Spain (n = 2; Cartwright and Colbourne, 2002; Galmes et al., 2003), Sweden (n = 1; Insulander et al., 2005), the United Kingdom (England and Wales [n = 49; [PHLS, 1995; PHLS, 1996b; PHLS, 1997a; PHLS, 1997b; PHLS, 1998; PHLS, 1999a; PHLS, 1999b; PHLS, 2000a;. PHLS, 2000c; PHLS, 2001a; PHLS, 2001b; PHLS, 2002a; PHLS, 2002b; PHLS, 2003; PHLS, 2004; PHLS, 2006a; PHLS, 2006b; Nichols et al., 2006], Scotland [n = 4; Smith-Palmer and Cowden, 2003, 2004; Smith-Palmer et al., 2005]) (Tables 12.2 and 12.3). These 136 outbreaks (91.2% [124] in disinfected water, 8.8% [12] in freshwater) have sickened 19,271 persons (95.8% [18,472] in disinfected water; 4.2% [799] in freshwater). This underscores Cryptosporidium’s emergence over the past two decades into a leading cause of recreationalwater-associated outbreaks of AGI. The average outbreak size was 142 persons (range 2 to 5449 persons); 149 persons for treated venues (range 3 to 5449), 67 people for untreated venues (range 2 to 418 persons). The potential for large-scale transmission is further highlighted with documentation of a single outbreak that sickened over 5000 people; 25 outbreaks infected more than 100 people in each, and four outbreaks infected over 1000 people each. Molecular typing (Xiao et al., 2004; Chapter 5) of specimens from 35 outbreaks detected 23 outbreaks caused by C. hominis, seven outbreaks caused by C. parvum, four outbreaks caused by both C. hominis and C. parvum, and one outbreak caused by both C. hominis and C. meleagridis (Tables 12.2 and 12.3). The difficulty in detecting outbreaks makes it likely that the number of reported outbreaks is less than what actually occured (Hellard et al., 2000).
A.
Outbreaks in the United States: General Overview
Since 1971, the U.S. Centers for Disease Control and Prevention (CDC), the U.S. Environmental Protection Agency (EPA), and the Council of State and Territorial Epidemiologists (CSTE) have maintained the Waterborne Disease and Outbreak Surveillance System (WBDOSS) that tracks the occurrences and causes of waterborne disease outbreaks (WBDOs) associated with drinking water. Tabulation of recreational water-associated outbreaks was added to the surveillance system in 1978. WBDOSS activities are intended to (1) characterize the epidemiology of WBDOs, (2) identify changing trends in the etiologic agents that caused WBDOs and determine why the outbreaks occurred, (3) encourage public health personnel to detect and investigate WBDOs, and (4) foster collaboration among local, state, federal, and international agencies on initiatives to prevent waterborne disease transmission. However, the WBDOSS has clear limitations. The data reported represent only a portion of the burden of illness associated with recreational water exposure in the United States; the surveillance information does not include endemic waterborne disease risks, nor are reliable estimates available for the number of unrecognized WBDOs and associated cases of illness. However, the number of reported recreational
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TABLE 12.2 Waterborne-Disease Outbreaks of Cryptosporidiosisa (n = 68) Associated with Recreational Water by Date, State, and Venue—United States, 1988–2004b State
Cases n = 14,679
Year
Month
2004 2004 2004
September August August
Illinois Colorado California
8 6d 336c
Pool Pool Pool
Type
Hotel Hotel Water park
Setting
Treated Treated Treated
Treatment
2004 2004
August July
Wisconsin Illinois
6 37
Community Community
Treated Treated
2004 2004
July June
Ohio Georgia
160d 14
Pool Pool/interactive fountain Pool Pool
Dziuban et al., 2006 Dziuban et al., 2006 Dziuban et al., 2006; Wheeler et al., 2007 Dziuban et al., 2006 Dziuban et al., 2006
References
Community Community
Treated Treated
Dziuban et al., 2006 Dziuban et al., 2006
2003 2003 2003 2003
August July July June
Arkansas Idaho Wisconsin Iowa
4 4 14 63
Swimming pool Lake Wading pool Wading pool
Treated Untreated Treated Treated
Dziuban et al., 2006 Dziuban et al., 2006 Dziuban et al., 2006 Dziuban et al., 2006
2003
June
Kansas
617d
Pool
Large facility Lake Community Daycare center Community
Treated
Dziuban et al., 2006
2002 2002 2002 2002
August August August July
Minnesota Minnesota Texas Georgia
16 41 54d 3
Pool Pool Pool Pool
Treated Treated Treated Treated
Yoder Yoder Yoder Yoder
2002
July
Massachusetts
767
Treated
Yoder et al., 2004
2002 2002 2002
July July May
Wisconsin Minnesota Wyoming
44 52 3
Pool/fountain/ whirlpool Lake Pool Lake
Resort Hotel Hotel Daycare center Health club State park Health club Lake
Untreated Treated Untreated
Yoder et al., 2004 Yoder et al., 2004 Yoder et al., 2004
2001
August
Wyoming
2
Hot spring water used in flow-through pool
State park
Untreated
Yoder et al., 2004
2001 2001
July July
Nebraska Illinois
21 358d
Pool Pool
Community Water park
Treated Treated
2001
July
Nebraska
157
Pool
Community
Treated
Yoder et al., 2004 Yoder et al., 2004; Causer et al., 2006 Yoder et al., 2004
2000 2000 2000
August August August
Florida Florida Colorado
19 5 112c
Pool Pool Pool
Treated Treated Treated
Lee et al., 2002 Lee et al., 2002 Lee et al., 2002
2000 2000
August August
Florida Minnesota
5 4
Pool Pool
Treated Treated
Lee et al., 2002 Lee et al., 2002
2000 2000 2000 2000 2000
July July July July July
Minnesota South Carolina Florida Minnesota Minnesota
6 26d 3 7 220d
Pool Pool Pool Pool/pond Lake
Resort Country club Municipal pool Condo Municipal pool Hotel Neighborhood Apartments Day camp Swimming beach
Treated Treated Treated Treated Untreated
Lee et al., 2002 Lee et al., 2002 Lee et al., 2002 Lee et al., 2002 Lee et al., 2002
et et et et
al., al., al., al.,
2004 2004 2004 2004
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TABLE 12.2 (CONTINUED) Waterborne-Disease Outbreaks of Cryptosporidiosisa (n = 68) Associated with Recreational Water by Date, State, and Venue—United States, 1988–2004b Year
Month
State
Cases N = 14,679
Type
Setting
Treatment
References
2000 2000 2000
July June June
Minnesota Georgia Ohio
15 36 700c,d
Lake Pool Pool
Beach Private pool Club
Untreated Treated Treated
Lee et al., 2002 Lee et al., 2002 CDC, 2001a; Lee et al., 2002; Mathieu et al., 2004 CDC, 2001a; Lee et al., 2002
2000
June
Nebraska
225d
Pools
Community
Treated
1999 1999
August August
Florida Florida
6 38
Private pool Beach
Treated Treated
1999 1999
July July
Minnesota Wisconsin
10 10
Pool Interactive fountain Pool Pool
Trailer park Municipal pool
Treated Treated
Lee et al., 2002 CDC, 2000; Lee et al., 2002 Lee et al., 2002 Lee et al., 2002
1998
August
Oregon
69
Pool
Treated
Barwick et al., 2000
1998 1998 1998 1998
July July July July
Wisconsin Pennsylvania Wisconsin Minnesota
9 8 12 7
Pool Lake Pool Pool
Treated Untreated Treated Treated
Barwick et al., 2000 Barwick et al., 2000 Barwick et al., 2000 Barwick et al., 2000
1998
July
Florida
7
Pool
Treated
Barwick et al., 2000
1998
June
Wisconsin
12
Pool
Treated
Barwick et al., 2000
1998
April
Minnesota
45
Pool
Community pool Public pool State park Public pool Community pool Daycare center Community pool Swim club
Treated
Barwick et al., 2000; Lim et al., 2004
1997
July
Minnesota
369c
Fountain
Zoo
Treated
CDC, 1998; Barwick et al., 2000
1996 1996 1996 1996
August August June June
Indiana California Florida Idaho
3 3000 77 55
Beach Water park Community Resort
Untreated Treated Treated Untreated
Levy et al., 1998 Levy et al., 1998 Levy et al., 1998 Levy et al., 1998
1996
June
Florida
22
Lake Pool Wading pool Hot spring water used in some water features Wading pool
Community
Treated
Levy et al., 1998
1995 1995
July July
Nebraska Georgia
14 5449d
Wading pool Pool
Water park Water park
Treated Treated
1995
June
Kansas
24
Pool
Swimming pool
Treated
Levy et al., 1998 Beach et al., 1996; Levy et al., 1998 Levy et al., 1998
1994
July
Missouri
101
Pool
Hotel
Treated
1994
July
New Jersey
418
Lake
State park
Untreated
1993
August
Wisconsin
64
Pool
Hotel
Treated
Wilberschied, 1995; Kramer et al., 1996 Kramer et al., 1996; Kramer et al., 1998 CDC, 1994; Kramer et al., 1996
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TABLE 12.2 (CONTINUED) Waterborne-Disease Outbreaks of Cryptosporidiosisa (n = 68) Associated with Recreational Water by Date, State, and Venue—United States, 1988–2004b Year
Month
State
Cases N = 14,679
Type
Setting
Treatment
References
1993
August
Wisconsin
54
Pool
Community
Treated
CDC, 1994; Kramer et al., 1996 Kramer et al., 1996 MacKenzie et al., 1995; Kramer et al., 1996
1993 1993
August April
Wisconsin Wisconsin
5 51
Pool Pool
Community Hotel
Treated Treated
1992 1992
August May
Idaho Oregon
26 500
Water slide Wave pool
Park Water park
Treated Treated
Moore et al., 1993 Moore et al., 1993; McAnulty et al., 1994
1988
August
California
44
Pool
Private pool
Treated
CDC, 1990, Sorvillo et al., 1992
Note: NA = not available. a
b c d
Five outbreaks identified other enteric pathogens in addition to Cryptosporidium. The cases attributed to these outbreaks have been classified as Cryptosporidium cases. 2004 is the last year of published summary data from the Waterborne Disease and Outbreak Surveillance System. Determined to be Cryptosporidium parvum by molecular methods (Chapter 5, Molecular Epidemiology). Determined to be Cryptosporidium hominis by molecular methods (Chapter 5, Molecular Epidemiology).
water-associated WBDOs of AGI have increased significantly since 1979 when CDC first began receiving AGI reports (Figure 12.4; Dziuban et al., 2006). The largest increases in number of outbreaks were in treated venues (Figure 12.5). These increases are likely to be due to a combination of factors that include the emergence of Cryptosporidium as a human pathogen, increased participation in aquatic activities (World Waterpark Association, 2006), increases in the number of aquatic venues (World Waterpark Association, 2006), and increased recognition, investigation, and reporting of recreational water-associated outbreaks. Cryptosporidium has emerged in the United States as the leading cause of recreational-water-associated outbreaks, particularly in artificial, treated venues. Since 1988, 68 cryptosporidiosis outbreaks associated with recreational water use have been reported to CDC (Table 12.2, CDC, 1990; Sorvillo et al., 1992; Herwaldt et al., 1991; Moore et al., 1993; Kramer et al., 1996; Levy et al., 1998; Barwick et al., 2000; Lee et al., 2002; Yoder et al., 2004; Dziuban et al., 2006; Craun et al., 2005). During 1988 to 2004, Cryptosporidium was the etiologic agent in 33.3% of reported AGI outbreaks; 7.4% of outbreaks associated with untreated, natural environments (lakes, rivers), and 59.6% of outbreaks associated with artificial, treated venues (e.g., swimming pools; Figure 12.6). During 2003 to 2004, the latest reporting data available, Cryptosporidium was the etiologic agent in 40% of reported outbreaks of AGI; 8.3% of outbreaks associated with untreated, natural environments, and 61.2% of outbreaks associated with artificial, treated venues (Dziuban et al., 2006). Cryptosporidium plays a larger role when the number of persons ill with AGI (14,679) is examined (Figure 12.7). Since 1988, cryptosporidiosis accounted for 70.7% of AGI cases reported to WBDOSS, 12.2% of cases associated with untreated, natural environments, and 91.6% of outbreaks associated with artificial, treated venues. Outbreaks have been identified in 22 states in the United States since 1988 (Figure 12.8). Treated venue outbreaks were documented from April to September and natural water outbreaks were documented from May through August (Figure 12.9). Only 14.7% (10/68) of the total cryptosporidiosis outbreaks were associated with use of natural waters, and all were associated with either lakes or hot spring water used for filling an artificial recreational water feature. The seasonality of cryptosporidiosis outbreaks reflects the seasonal pattern of late summer to early fall cryptosporidiosis case reporting in the United States (Figure 12.2). Molecular typing (Xiao et al., 2004; Chapter 5) of specimens from 13 U.S. outbreaks detected nine outbreaks caused by C. hominis, three outbreaks caused by C. parvum, and one outbreak caused by both C. hominis and C. parvum (Table 12.2).
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TABLE 12.3 Waterborne Disease Outbreaks of Cryptosporidiosisa (n = 68) Associated with Recreational Water by Date, Country, and Venue—International Sites Excluding the United States, 1988–2005b Year
Month
Country
Cases (n = 4592)
Type
Setting
Treatment
2005
April
Australia
>16
Pools (n = 4)
Four separate facilities
Treated
1998 1997
February December
Australia Australia
125 >1000
1997 1997 1994
December November November
Australia Australia Australia
>15 >18 >17
Pools (n = 7) Pools, person to person Pool Two pools Indoor pool
Community Treated Statewide Treated outbreak Pool complex Treated Community Treated Community Treated outbreak
2003
July
Canada
33
Pool complex
2001
May
Canada
59
Pool
1990
October
Canada
89
Children’s pool
Recreational complex Hotel, dance festival Recreational complex
2005
August
Japan
38d
Pool
2004
August
Japan
222
1998
February
New Zealand
2003 2000
June July
2002
Reference Paterson and Goldthorpe, 2006; Black and McAnulty, 2006 Hellard et al., 2000 Puech et al., 2001 Stafford et al., 2000 Beers et al., 1998 Lemmon et al., 1996
Treated
Louie et al., 2004
Treated
Macey et al., 2002
Treated
Bell et al., 1993
Hotel
Treated
Pool
Hotel
Treated
Yokoi et al., 2005 Ichinohe et al., 2005
53
Pool
Community pool
Treated
Baker et al., 1998
Spainf Spainf
391c 172d
Pool Pool
Hotel Hotel
Treated Treated
Galmes et al., 2003 Cartwright et al., 2002; PHLS, 2000b
August
Sweden
1000c
Pool/children’s pool
Community
Treated
Insulander et al., 2005
2005 2005 2005 2005
September August July May
United Kingdom England/Wales England/Wales England/Wales England/Wales
>129 88d 15 3
Pools Pools Paddling pool Pool
Treated Treated Treated Treated
PHLS, 2006b PHLS, 2006b PHLS, 2006b PHLS, 2006b
2004
October
England/Wales
6d
Pool
Treated
PHLS, 2006a
2004
October
England/Wales
10d
Pool
Treated
PHLS, 2006a
2004 2004 2004 2003 2003
October May March October August
England/Wales England/Wales England/Wales England/Wales England/Wales
12 7d 13 2 122de
Pool Pool Pool Pool Interactive water feature
N/A N/A Wildlife park Leisure center Leisure center Leisure center School Public pool Public pool Private pool Public park
Treated Treated Treated Treated Treated
PHLS, 2006a PHLS, 2006a PHLS, 2006a Nichols et al., 2006 PHLS, 2004
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TABLE 12.3 (CONTINUED) Waterborne Disease Outbreaks of Cryptosporidiosisa (n = 68) Associated with Recreational Water by Date, Country, and Venue—International Sites Excluding the United States, 1988–2005b Year
Month
Country
Cases (n = 4592)
Type
Setting
Ornamental fountain Pool Interactive water feature Pool Hydro training pool Pool
Water feature complex Sports center Leisure facility Public pool N/A
2003
August
England/Wales
4
2003 2003
August August
England/Wales England/Wales
17d 63cd
2003 2003
August January
England/Wales England/Wales
22 66d
2002
September
England/Wales
20cd
2002
April
England/Wales
4
Various water sources
2001
October
England/Wales
3d
Pool
2001
August
England/Wales
14
2001
June
England/Wales
2001
February
2000
Treatment
Reference
Treated
PHLS, 2004
Treated Treated Treated Treated
PHLS, 2004 PHLS, 2004; Jones et al, 2006 PHLS, 2004 PHLS, 2003a, 2004
Treated
PHLS, 2003
Treated
PHLS, 2003
Treated
PHLS, 2002a, 2001b PHLS, 2002a, 2001b PHLS, 2001b, 2002b PHLS, 2001b, 2002b PHLS, 2001a, 2001b PHLS, 2001a, 2001b PHLS, 2001a, 2001b PHLS, 2001a, 2001b PHLS, 2001a, 2001b PHLS, 2001a, 2001b PHLS, 2000c, 2001b PHLS, 2000a, 2000c PHLS, 2000c
Stream
Leisure complex Water activity center Leisure complex Beach
Untreated
152d
Pool
School
Treated
England/Wales
5
Pools
Public pool
Treated
October
England/Wales
5cd
Pool
Club
Treated
2000
September
England/Wales
9
Pool
Public pool
Treated
2000
September
England/Wales
12c
Pool
Public pool
Treated
2000
September
England/Wales
10d
Pool
Public pool
Treated
2000
September
England/Wales
7
Pool
Public pool
Treated
2000
August
England/Wales
3d
Pool
Public pool
Treated
2000
May
England/Wales
41c
Pool
Treated
1999
November
England/Wales
19
Pool
1999
September
England/Wales
30
Pools
1999
September
England/Wales
8
Pool
1999
August
England/Wales
54
Pool
Leisure complex Leisure complex Leisure complex Leisure complex N/A
Treated
1999
August
England/Wales
16d
Pool
N/A
Treated
1999
July
England/Wales
14
Pool
Treated
1999
July
England/Wales
11
Pool
Leisure complex N/A
Treated
1998
November
England/Wales
9
Pool
Public pool
Treated
1998
September
England/Wales
14d
Pool
Swimming complex
Treated
Treated Treated Treated
PHLS, 2000a, 2000c PHLS, 2000a, 2000c PHLS, 2000a, 2000c PHLS, 2000a, 2000c PHLS, 2000a, 2000c PHLS, 1999a, 1999b PHLS, 1999a, 1999b
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TABLE 12.3 (CONTINUED) Waterborne Disease Outbreaks of Cryptosporidiosisa (n = 68) Associated with Recreational Water by Date, Country, and Venue—International Sites Excluding the United States, 1988–2005b Year
Month
Country
Cases (n = 4592)
Treatment
Reference
1998 1997 1997 1996
March May May July
England/Wales England/Wales England/Wales England/Wales
6 9 13 8
Pool Learner pool River Pool
Type
Fitness club N/A River N/A
Setting
Treated Treated Untreated Treated
1995
August
England/Wales
3
Pool
Treated
1994 1994 1994 1993
October N/A N/A January
England/Wales England/Wales England/Wales England/Wales
14 4 3 23
Pool Pool Pool Pool
Treated Treated Treated Treated
PHLS, 1995 Nichols et al., 2006 Nichols et al., 2006 Furtado et al., 1998
1992 1988 2004
March August N/A
England/Wales England/Wales Scotland
13 67 6
Paddling pool Learner pool Pool
Leisure center School Public pool Country club Leisure center Community Sports center N/A
PHLS, 1998, 1999b PHLS, 1997b PHLS, 1997b PHLS, 1997a; Sundkvist et al., 1997 PHLS, 1996b
Treated Treated Treated
2003
N/A
Scotland
10
Pool
N/A
Treated
2003
N/A
Scotland
51
Pool
Public baths
Treated
2002
N/A
Scotland
75
Pool
Leisure pool
Treated
Hunt et al., 1994 Joce et al., 1991 Smith-Palmer et al., 2005 Smith-Palmer and Cowden, 2004 Smith-Palmer and Cowden, 2004 Smith-Palmer and Cowden, 2003
Note: NA = not available. a b c d e f
Some outbreaks may have identified other enteric pathogens in addition to Cryptosporidium. Some counties have not yet published summaries of outbreaks through 2005. Determined to be Cryptosporidium parvum by molecular methods; Nichols et al., 2006. Determined to be Cryptosporidium hominis by molecular methods; Nichols et al., 2006. Determined to be Cryptosporidium meleagridis by molecular methods; Nichols et al., 2006. All returning U.K. tourists.
Number of outbreaks
25 20 15 10 5
19 78 19 80 19 82 19 84 19 86 19 88 19 90 19 92 19 94 19 96 19 98 20 00 20 02 20 04
0
Year FIGURE 12.4 Number of recreational water-associated outbreaks of acute gastroenteritis (n = 207) by year—United States, 1978–2004.
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16 Treated
Number of outbreaks
14
Untreated
12 10 8 6 4 2
20 04
20 02
20 00
19 98
19 96
19 92 19 94
19 88 19 90
18 94 19 86
19 82
19 80
19 78
0
Year FIGURE 12.5 Number of recreational water-associated outbreaks of gastroenteritis (n = 207), by water type and year—United States, 1978–2004.
Type of Exposure (n=189)
Etiologic Agent (n=189) Norovirus 11.1%
Treated water 49.7%
Cryptosporidium spp. 33.3%
Shigella spp. 15.9% E. coli 10.6%
Untreated water 50.3%
Giardia spp. 7.4%
Otherb 4.2%
Unidentified 17.5%
Etiologic Agent: Untreated Water (n=95) Shigella spp. 24.2%
b
E. coli 16.8%
Cryptosporidium spp. 59.6%
Norovirus 13.7%
Other 3.2%
Otherb 5.3% Cryptosporidium spp. 7.4%
Unidentified 27.4%
Etiologic Agent: Treated Water (n=94)
Giardia spp. 7.4%
Unidentified 7.4% Shigella sonnei 7.4% Giardia intestinalis 7.4%
Norovirus 8.5% E. coli 4.3%
FIGURE 12.6 (A color version of this figure follows page 242.) Recreational water-associated outbreaks of acute gastroenteritis by type of exposure and etiologic agent—United States, 1988–2004a. aFive outbreaks involved Cryptosporidium with one or more other pathogens detected in greater numbers; these outbreaks have been classified as non-Cryptosporidium outbreaks; bThese include outbreaks of Campylobacter, Plesiomonas, Salmonella, and suspected chemical or toxin etiologies.
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Type of Exposure (n=20,441)
Etiologic Agent (n=20,441)
Treated water 73.7%
Cryptosporidium spp. 70.7%
Unidentified/ Otherb 11.2% Giardia spp. 2.3%
Untreated water 26.3%
E. coli 2.3% Shigella spp. 9.4%
Etiologic Agent: Untreated Water (n=5,386) Shigella spp. 31.5%
Etiologic Agent: Treated Water (n=15,055)
E. coli 7.7%
Cryptosporidium spp. 91.6%
Norovirus 9.5%
Unidentified/ Otherb 37.2%
Norovirus 4.3%
Unidentified/ Otherb 1.7% Shigella sonnei 1.5% Cryptosporidium Giardia intestinalis 2.5% spp. 12.2% E. coli 0.4% Norovirus 2.4% Giardia spp. 1.7%
FIGURE 12.7 (A color version of this figure follows page 242.) Recreational water-associated cases of acute gastroenteritis by type of exposure and etiologic agent—United States, 1988–2004a. aFive outbreaks involved Cryptosporidium with one or more other pathogens detected in greater numbers; the cases attributed to these outbreaks have been classified as nonCryptosporidium cases; bOther includes outbreaks of Campylobacter, Plesiomonas, Salmonella, Cyanobacter, and suspected chemical etiologies.
4+ (five states)† 3 (three states) 2 (four states) 1 (10 states) 0 (28 states)
FIGURE 12.8 Number of recreational water-associated outbreaks of cryptosporidiosis (n = 68)—United States, 1988–2004*. *Note that these numbers are largely dependent on reporting and surveillance activities in individual states, and do not necessarily indicate the true incidence in a given state; †Florida: nine outbreaks; Georgia: four outbreaks; Minnesota: 12 outbreaks; Nebraska: four outbreaks; Wisconsin: 11 outbreaks.
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Untreated Venues
30
Treated Venues
No. of outbreaks
25 20 15 10 5 0 J
F
M
A
M
J
J
A
S
O
N
D
Month FIGURE 12.9 Number of recreational water-associated outbreaks of cryptosporidiosis (n = 68) by venue and month— United States, 1988–2004.
B.
Outbreaks in Countries Other than the United States: General Overview
Since 1988, 68 other reported outbreaks of cryptosporidiosis from seven countries have been associated with recreational water (Table 12.3). Of these outbreaks, 97.1% (67/69) were associated with treated water such as swimming pools or interactive fountains. At least 4,592 persons were sickened with 99.4% (4,565) of illnesses associated with treated water venues. England and Wales introduced enhanced surveillance for infectious intestinal disease in 1992 after a pilot study to classify and standardize the approach for data collection and outbreak classification (Nazareth et al., 1994). A more extensive list of criteria was developed to categorize the levels of evidence needed for designating outbreaks as waterborne (PHLS, 1996a; Tillett et al., 1998). From 1992 to 2005, there have been at least 49 outbreaks of cryptosporidiosis, with 95.9% (47/49) in treated pools or fountains (Table 12.3). From 1992 to 2003, the parasite was associated with 95% (38/40) of all recreational-waterassociated outbreaks in England and Wales (Smith et al., 2006). The seasonality for these outbreaks is similar to those in the United States with a peak in the late summer to early fall months of August through October: a marked contrast with the spring seasonal occurrence for drinking-water-associated outbreaks (Smith et al., 2006). Molecular typing (Xiao et al., 2004; Chapter 5) of specimens from 22 outbreaks outside the United States detected 14 outbreaks caused by C. hominis, four outbreaks caused by C. parvum, three outbreaks caused by both C. hominis and C. parvum, and one outbreak caused by both C. hominis and C. meleagridis (Table 12.3).
C. 1.
Recreational Water Types and Outbreak-Associated Deficiencies Marine Water
No outbreaks of cryptosporidiosis have been associated with marine beach use. However, outbreaks of AGI or cryptosporidiosis associated with swimming at large beaches that draw from a wide geographic range might be difficult to detect because potentially infected persons disperse widely from the site of exposure and, therefore, might be less likely to be identified as part of an outbreak. Fecal contamination of marine beaches is well documented. From 1997 to 2005, 21 to 28% of the reporting coastal and Great Lakes beaches in the United States were annually impacted by beach closures or advisories related to microbial water quality indicators (i.e., detection of fecal contamination) that exceeded EPA standards. For the 2005 swimming season, there were closures on 4% (27,177/743,036) of beach days at the 4,025 monitored beaches in the United States (EPA, 2006). Despite documentation of fecal contamination at
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these beaches, the U.S. WBDOSS has not reported an ocean-associated outbreak of AGI or cryptosporidiosis since 1978. In addition, no marine water-associated outbreaks of cryptosporidiosis have been reported from other countries. However, prospective epidemiologic studies in the United States and elsewhere (Stevenson, 1953; Cabelli et al., 1979; Dufour, 1984; Seyfried et al., 1985; Cheung et al., 1990; Ferley et al., 1989; Fleisher et al., 1998; Pruss, 1998; Haile et al., 1999; Henrickson et al., 2001; Wade et al., 2006; Weidenmann et al., 2006) have clearly demonstrated that ocean beaches and large bodies of freshwater (e.g., the Great Lakes in the United States; Wade et al., 2006) can be a significant source of AGI in swimmers. Undetected transmission of cryptosporidiosis may be occurring based on findings that Cryptosporidium can survive for long periods of time in seawater and remain infectious in shellfish (Fayer et al., 1998a; Robertson et al., 1992; Tamburrini and Pozio, 1999).
2.
Freshwater Lakes, Rivers, and Hot Springs
Despite evidence of ubiquitous contamination of freshwater lakes and rivers by Cryptosporidium and quantitative data suggesting that Cryptosporidium concentrations were, in some cases, high enough to transmit the parasite (LeChevallier et al., 1991; LeChevallier and Norton, 1995; Xiao et al., 2001; Ward et al., 2002; Montemayor et al., 2005; Coupe et al., 2006; Karanis et al., 2006), only 12 outbreaks (ten in the United States and two in England and Wales) have been associated with freshwater use (Tables 12.2 and 12.3). Only 8.8% (12/136) of cryptosporidiosis outbreaks have been associated with freshwater use. Locations have included eight lakes, two rivers/streams, and two uses of natural hot spring water in artificial recreational venues with no added disinfection or treatment. This low number of outbreaks may reflect poor detection or reporting of such outbreaks. It may also reflect that contamination is sporadic and that higher concentrations of Cryptosporidium are only transient after environmental events (e.g., heavy rainfall). In addition, natural recreational water areas may be aided by dilution of incoming runoff as well as natural circulation and bioremediation of the recreational water. These dilution effects may substantially reduce the risk of transmission in natural waters when compared to the smaller volumes found in treated venues. The lower mean size of the 12 documented freshwater outbreaks (67 persons versus 149 persons for treated venues) may reflect a lower concentration of pathogen, a lower bather load in freshwater venues, or increased challenges in detecting and investigating freshwater-associated outbreaks. In 1994, over 400 people contracted cryptosporidiosis when a lake in New Jersey (Kramer et al., 1998) had a pump failure that released sewage onto the ground near a recreational lake. Subsequent heavy rainfall likely washed the sewage into the lake. Water testing detected levels of fecal coliforms (500 and 1600 cfu/100 mL) exceeding the state standard of 200 cfu/100 mL. Contributing factors to the month-long outbreak included the small size of the lake and swimming beach and high levels of fecal indicator bacteria. In 2000, a lake-associated outbreak (Minnesota; Lee et al., 2002) affected 220 persons. No sources of contamination such as contaminated runoff, or broken septic or sewer lines were identified. However, the beach was heavily used by young children and molecular testing demonstrated seven fecal samples contained C. hominis indicating bather contamination as the source. In 2002 an outbreak of cryptosporidiosis in Wisconsin (Yoder et al., 2004) was associated with use of a swimming beach on one of the Great Lakes; the only Great Lakes outbreak documented since reporting started in 1978. The outbreak sickened at least 44 people with “sewage poisoning”; pathogens included Cryptosporidium, norovirus, and Shigella. The environmental investigation revealed high E. coli levels (> 2400 cfu/100 mL) on multiple occasions after the outbreak. Contamination might have resulted from bathers or dumping of sewage from boats moored offshore. After the investigation, beachclosure policies were changed, and the beach was subsequently closed to swimming five more times that summer; in each instance closure was <48 h after a rainfall event. These outbreaks support the use of routine fecal indicator testing of natural waters, particularly when new rapid tests are available that allow beach managers to obtain results and make closure decisions within several hours. However, freshwater beaches at small lakes with shallow swimming areas that have low or poor circulation may be particularly vulnerable to contamination by bathers; such contamination may not be routinely detected by traditional fecal indicator testing.
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Only two outbreaks have been associated with rivers or streams. In 2001, recreational exposure to stream water that flowed through a swimming beach in the United Kingdom caused 14 cases of gastroenteritis (5 patients were <15 years of age). Possible sources of contamination included release of treated wastewater effluent and illegal disposal of waste from campsite chemical toilets into a sewer. High levels of fecal coliforms and fecal streptococci were documented (PHLS, 2001b, 2002a). In 1997, 13 people in the United Kingdom became ill with AGI (seven confirmed cryptosporidiosis, one confirmed Giardia intestinalis) after visiting an activity center where they swam in a river after a heavy rainfall. Sources may have included runoff from nearby fields that were using a slurry of animal feces for fertilization (PHLS, 1997b). In two outbreaks in the United States, natural mineral or hot spring water were used to fill flowthrough pools or recreational water features at a resort water park (Levy et al., 1998; Yoder et al., 2004). The use of mineral or hot spring water is unlikely to affect the viability of enteric pathogens such as Cryptosporidium. As a result, transitioning the use of this untreated water from the traditional soaking to full body recreation in an enclosed artificial structure without routine disinfection and filtration would be expected to compromise water quality and increase the risk of transmission for all enteric pathogens, not just Cryptosporidium. Lessons learned from these freshwater-associated outbreaks that may reduce risk include: (1) routine fecal indicator monitoring would likely have closed the venues to swimming in some of these outbreaks, (2) routine sanitary surveys would likely have identified problems (e.g., leaking septic tanks, illegal dumping of waste) that could have been corrected before an outbreak, (3) swimming after heavy rainfall may increase risk of transmission, (4) dumping of human waste from boats may contribute to risk on nearby swimming beaches, (5) use of animal waste on watershed land near bathing beaches needs to be monitored, and (6) using untreated mineral water in artificial, full-body recreational pools increases the risk for enteric disease transmission compared to traditional use of this water for soaking activities.
3.
Disinfected Water
Over 90% (124/136) of reported recreational-water-associated outbreaks of cryptosporidiosis have been associated with swimming pools, and spray features such as interactive fountains (Tables 12.1 and 12.2). The circumscribed volumes of water, high bather loads, heavy use by diaper- and toddler-aged children, and the parasite’s chlorine resistance undoubtedly increase the potential for contamination and efficient transmission. The modes of contamination seem clear; fecal contamination by fellow swimmers such as diaper- and toddler-aged children. Only one of the 125 outbreaks (Joce et al., 1991) has been associated with pool plumbing problems that allowed raw sewage to enter the pool.
a.
Swimming Pools, Water Parks
The potential for massive transmission of the parasite is illustrated by water parks, which have been increasing in number and attendance over the past decade (World Waterpark Association, 2006); water park outbreaks have made 500 people ill in 1992 (McAnulty et al., 1994), 5449 persons in 1995 (Levy et al., 1998), 3000 persons in 1996 (Levy et al., 1998), 358 persons in 2001 (Causer et al., 2006), and 336 persons in 2004 (Wheeler et al., 2007). In addition, as with other enteric pathogens, cryptosporidiosis has the potential for transmission and amplification via multiple vehicles and routes. In at least two instances, recreational water played an amplifying role in outbreaks that expanded into community(Dziuban et al, 2006) or statewide (Puech et al., 2001) outbreaks. In 2003, an increased incidence of laboratory-confirmed cryptosporidiosis led to identification of an ongoing communitywide outbreak with multiple clusters (Kansas; Dziuban et al., 2006). The outbreak lasted for weeks and although the initial point of transmission could not be identified, transmission was associated with swimming pools used by swim teams and day camps, as well as multiple day care facilities where water-related activities were prevalent. It was clear that ill pool patrons and day care center attendees continued swimming despite their illness and exposed large numbers of persons to Cryptosporidium. In contrast, the next year there was a Cryptosporidium outbreak (Ohio; Dziuban et al., 2006) associated with a community swimming pool. The outbreak demonstrated that a rapid communitywide public health response during the early stages of an outbreak may help control the potential
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spread of enteric illness in a community. In Ohio, detection and investigation started within 1 week of exposure. The response included mitigating actions (e.g., hyperchlorination of all community pools and providing instructions regarding proper water hygiene to pool staff and users, day care centers, restaurants, and other potentially affected facilities). Hyperchlorination of the pool appeared to be effective in stopping transmission, and a rapid response to the outbreak was likely responsible for preventing further spread. As a result, the investigation detected no transmission outside of the single community pool, in contrast to the outbreak in Kansas. An increase in statewide reports of cryptosporidiosis during 1997 in New South Wales, Australia (Puech et al., 2001) led to the finding of a prolonged, statewide outbreak of cryptosporidiosis that affected over 1000 persons and was associated with swimming in public pools. Control measures included media statements for the public (e.g., do not swim when ill with diarrhea), advice to pool operators (e.g., prompt removal of fecal material from pools, improved pool maintenance), and creation of health communication materials and protocols for minimizing contamination of pools and spas. These community outbreaks illustrate that preventing amplification and expansion of outbreaks needs a rapid public health response and increased community involvement. In 1995 the largest documented (5449 persons ill) outbreak of cryptosporidiosis related to recreational water was in Georgia (Beach et al., 1996; Levy et al., 1998). The investigation determined that a water park was involved and that the risk factors for cryptosporidiosis were swallowing water and using the children’s pools. Both stool exams and water analyses demonstrated the presence of C. hominis and Giardia intestinalis. The main children’s pool was cofiltered with other features in the water park, which contributed to further transmission via cross-contamination of these other water features. Another water park outbreak (Illinois) sickened >358 persons and included cofiltered pools (Causer et al., 2006). The highest risk of cryptosporidiosis was associated with using the three pools (including a toddler pool) included on one filtration system. Cofiltered pools also were contributing factors in outbreaks in New Zealand (Baker et al., 1998) and in the United Kingdom (Hunt at al., 1994). Replenishing of water in a learner’s pool with water from the main adult pool may have played a role in transmission in another pool-associated outbreak in the United Kingdom (Sundkvist et al., 1997). An alternative pool design was found to be implicated in an Australian outbreak. The design included a 25-m swim area, a children’s learning area, and a bubble area popular with children all in the same pool with a single filtration system (Lemmon et al., 1996). These features warrant design changes. Most important would be separation of filtration systems in these facilities to prevent cross-contamination of cofiltered water features, particularly high-risk areas such as children’s pools. In August 2000, public health officials detected an increase in the number of cases of cryptosporidiosis over the baseline from the previous year (Nebraska; CDC, 2001a). Most of the initial cases were linked to swimming in two private swimming facilities. The pools at both locations were closed for 2 weeks and hyperchlorinated. Although the appropriate measures to disinfect the water were taken by staff, the outbreak expanded to other pools. Surveys indicated that 18% of respondents reported swimming while ill and 32% swam while ill or during the 2 weeks after resolution of symptoms, despite the pool’s closure. This was accomplished by swimmers moving to other community swimming pools. As a result, the outbreak expanded to include other pools and prolonged the outbreak for the entire summer until the end of the outdoor swimming season. This outbreak underscores the need to respond differently to outbreaks associated with recreational water compared to some foodborne outbreaks. In restaurantassociated foodborne outbreaks, the restaurant is closed until the cause of the outbreak is identified. Although this may halt the spread of foodborne outbreaks, the same principle is less effective in curtailing swimming pool-associated diarrheal outbreaks as pool closure results in infected persons moving to new swimming sites. To decrease this chance, it is advisable to remediate the pool as quickly as possible (i.e., overnight hyperchlorination) and reopen rapidly. In August 2000, another outbreak affected more than 700 persons in Ohio (CDC, 2001a; Mathieu et al., 2004). The outbreak, which was associated with swimming in a local pool, started in late June and continued through September. The prolonged nature of the outbreak suggests that people continued swimming while they were still ill; at least four formed fecal accidents and one diarrheic fecal accident were documented during this time. In addition, laboratory testing confirmed C. hominis and C. parvum in patient stools suggesting that there were at least two separate contamination events. Enhanced communication between health personnel, pool operators, and
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swimmers is essential to alert swimmers about outbreaks and decrease the chance of ill swimmers recontaminating swimming facilities. Periodic hyperchlorination or “shock” treatments should also be utilized to keep potential oocyst levels down. Remediation of pools following fecal accidents has been accomplished in multiple ways depending on whether the feces released is formed or diarrheic. Formed stools have been treated as either noninfectious events (WHO, 2006) in which case the formed feces is removed and discarded but the pool remains open, or as potentially infectious contaminants. The U.S. CDC tested 293 formed fecal accident stools collected from pools and found 4.4% positive for Giardia and none positive for Cryptosporidium. Based on these data, CDC guidelines recommend that the pool be closed, the formed stool be removed, and sufficient time be allowed to pass for inactivation of the moderately chlorine-resistant cysts of Giardia (e.g., 1 ppm free chlorine for 30 min; CDC, 2001c). With diarrheic stools, the potential for transmission of infectious agents is increased, and the probability of the diarrhea containing Cryptosporidium, although unknown, would be expected to be higher than with formed stool. When outbreaks of cryptosporidiosis began to increase in the 1990s, some decontamination protocols included draining the pool, steam cleaning or disinfecting the surfaces, replacing or steam cleaning filter media, and refilling the pool (CDC, 1990; Joce et al., 1991; Bell et al., 1993). No supportive data were collected to demonstrate that this was necessary to reduce transmission, particularly in light of the length of time for the process and the common occurrence of fecal accidents in the swimming environment. Current practices generally recommend that swimming pools be hyperchlorinated to levels that ensure attainment of the 9600 CT value for Cryptosporidium (e.g., 8 h at 20 ppm free chlorine; Korich et al., 1990; Kebabjian, 1995; CDC, 2001c; WHO, 2006) or allow multiple pool volumes/turnovers (e.g., six) to pass through the filtration system (WHO, 2006). In 2004 an outbreak associated with a small water park in California affected >250 persons (Wheeler et al., 2007). Of employees interviewed, 83% were ill with diarrhea and the peak of illness in employees preceded the peak for patrons, suggesting contamination by employees contributed to the outbreak. Upon interview, three employees and 16 patrons admitted swimming while ill with diarrhea. Employees were required to be in the pool each day as part of their job. Other outbreaks have also documented illness in employees including the first documented outbreak of recreational-water-associated cryptosporidiosis (Sorvillo et al., 1992). During this outbreak in California, 75% (3/4) of lifeguards were ill, along with patrons, in a dose-dependent manner (length of time in pool). An outbreak in 1992 in British Columbia, Canada, 27% (Bell et al. 1993) of employees were ill and another outbreak investigation documented employees working while they were ill (Louie et al., 2004). None of these three investigations documented onsets of illness relative to patrons, so it was not possible to ascertain if employees played a role in initial amplification of the parasite. Pool operators and public health officials need to be sensitive to reports of diarrhea in aquatics staff because they may be sentinels for potential outbreaks of swimmingassociated cryptosporidiosis. Diarrhea exclusion policies need to be developed for all aquatics facilities. Employee policies which do not include sick leave need to find alternative non-water-related work for ill employees to prevent employees from continuing to work while ill with diarrhea. Because traditional chlorination has little effect on Cryptosporidium, the traditional use of tap water to fill or operate temporary aquatic venues (e.g., wading pools, paddling pools, and waterslides) used by young children also needs to be reconsidered, particularly in institutional settings (e.g., day care centers and schools). If the water is not treated with adequate levels of disinfectant, residual disinfectant in the water is rapidly depleted; users are then at higher risk of exposure to infectious microbes in the untreated water. Special consideration needs to be given to kiddie pools, some of which have had unfavorable water-quality test results and associations with previous outbreaks (CDC. 2001d; PHLS, 2001b). In one Cryptosporidium outbreak (Iowa; Dziuban et al., 2006), a kiddie pool at a day care facility was filled with potable municipal water that had not received additional treatment, expediting infection of children and eventual expansion into a communitywide outbreak. In Kansas (Dziuban et al., 2006), the communitywide outbreak also involved use of kiddie wading pools and in-ground pools at local day care centers. Portable waterslides in which municipal water is used might be overlooked as sources of disease transmission because they can be set up, used, and taken down in a matter of hours. As a result, the use of temporary pools filled with municipal water that do not include routine disinfection
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and filtration should be considered carefully by the public and, based on documented outbreaks, should be eliminated from institutional settings (e.g., day care centers and schools; Kirkpatrick et al., 2006). Travel of infected persons during outbreaks may expedite the spread of cryptosporidiosis. The largest waterborne outbreak of cryptosporidiosis ever recorded began in March 1993 (MacKenzie et al., 1994). The outbreak was associated with drinking water from the Milwaukee, WI, municipal drinking water system. In April, 1993, an outbreak of cryptosporidiosis was associated with a hotel in a community approximately 70 miles from Milwaukee (MacKenzie et al., 1995). It was demonstrated that the main risk factor for illness was swimming in the hotel pool. Although not proved, the authors felt the most plausible explanation was that residents of Milwaukee that were ill with cryptosporidiosis (Milwaukee residents were known to have stayed at the hotel) contaminated the pool. The outbreak underscored the potential for amplifying cryptosporidiosis through swimming activities during an ongoing outbreak. It also highlighted the need to educate the general public about the need to refrain from swimming while ill with diarrhea during outbreaks. By April, diarrhea rates had fallen significantly in Milwaukee, so a specific message about swimming was not initiated after this satellite outbreak came to light (MacKenzie et al., 1995). Similarly, in 2000, a swim-club-associated outbreak involved 700 persons in Ohio (Lee et al., 2002). A family from Ohio who were members of the implicated swim club vacationed in Florida after their child became ill. The ill infant was allowed to continue swimming, had two fecal accidents in the pool, and at least five people became ill following use of the pool. International travel as a risk factor for cryptosporidiosis has also been documented (Roy et al., 2004; Hunter et al., 2004), but associated activities during international travel have not been well delineated. Two outbreaks in Spain in 2000 and 2003 illustrate one of these risk factors (Cartwright and Colbourne, 2002; Galmes et al., 2003). Both outbreaks involved tourists returning from vacations in Spain that were detected by surveillance systems in the United Kingdom. In 2001, 22 confirmed cases of cryptosporidiosis were reported through surveillance systems in Scotland and England and Wales. All persons had been on holiday at the same hotel in Spain, and Cryptosporidium was detected in the hotel’s swimming pool filters (Cartwright and Colbourne, 2002). In 2003, another outbreak was detected by Scottish public health authorities that involved tourists returning from holiday in Spain. At least 391 tourists from England, Wales, Scotland, and Northern Ireland were affected by the outbreak (Galmes et al., 2003). The hotel swimming pool was implicated, and Cryptosporidium was found in the pool water. Travelassociated cryptosporidiosis may be difficult to detect because of the dispersion of persons from the exposure site to widely scattered locations. However, when groups travel together and meet regularly after returning from travel, they may be more likely to discuss illness that may be reported to public health authorities and serve as sentinels for travel-associated disease transmission. Such was the case with cryptosporidiosis outbreaks associated with a dance event in Canada (Macey et al., 2002) and two swim team events in Japan (Ichinohe et al., 2005; Yokoi et al., 2005). Other recreational water outbreaks associated with events such as sports events or other team-related activities have been documented (Dziuban et al., 2006). These outbreaks highlight the difficulty with detecting outbreaks that involve more than one country and the need for improved operation of recreational water facilities on a global basis. It also is important to stress education of travelers about infectious disease transmission associated with recreational water use.
b.
Interactive Fountains/Splash Pads/Spray Features
At least seven outbreaks of cryptosporidiosis have involved interactive fountains or water spray features that are either intended for, or accessible to, recreational use (Table 12.2 and 12.3). These attractions do not usually have standing water as part of the design. However, they usually have recirculating water and may also have some level of secondary water treatment (e.g., filtration, disinfection). Because these water features do not have standing water, they may not fall under existing pool regulations that were designed to cover the traditional water-filled swimming pool. The first documented outbreak was associated with a decorative fountain at a zoo in Minnesota (CDC, 1998). Spray nozzles located beneath ground level metal grates pumped jets of water up through the grating to a height of 1 to 6 ft. The water returned through a chlorinator and filter before being recirculated. Although the fountain was not intended for public immersion, investigators observed that children would regularly stand over the nozzles and let the water jets soak their entire bodies. Cryptosporidium parvum
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was identified as the cause of the outbreak. Following the outbreak, the zoo erected a fence around the entire fountain to prevent entry into the water jets. The second recorded outbreak of AGI associated with an interactive fountain associated outbreak of AGI (Florida; CDC, 2000) was caused by Cryptosporidium and Shigella. The presence of the chlorine-sensitive Shigella indicated that the facility maintenance was deficient. The fountain had been open less than a month and used recirculated water that drained into a reservoir where the water was chlorinated but not filtered. Chlorine tablets that had filled the chlorinator were depleted and had not been replaced since the opening of the facility. As mentioned earlier, diaperand toddler-aged children were observed standing over the water jets and soaking their entire bodies. A filtration system and chlorine monitor was installed, signs were posted to discourage water swallowing and encourage showers before entry, and diaper-aged children were excluded. Improved maintenance of all aquatic facilities is needed to prevent transmission of chlorine-sensitive pathogens such as Shigella. Three fountain-associated outbreaks were detected in 2003 in the United Kingdom (one was an ornamental fountain), further highlighting the need for attraction guidelines (PHLS, 2004; Jones et al., 2006). One outbreak sickened 63 children and was characterized by a high attack rate (83%), long duration of illness (median 8 days), and relatively severe disease (16% hospitalization rate). The disinfection and filtration systems were improved and new policies (e.g., removal of footwear before entering) were instituted (Jones et al., 2006). Another outbreak in 2003 involved an interactive fountain where 122 persons became ill, over 80% of whom were under 15 years of age (Jones et al, 2006). The third outbreak in 2003 involved a decorative fountain that children used for play. The large body of enclosed water was used as a paddling pool despite the decorative intent of the fountain (Jones et al., 2006). The largest outbreak to date that has been associated with interactive fountains was in New York in 2006 (Schaffzin et al., 2006). Several thousand persons became infected with C. hominis following use of a spray park. The filtration and disinfection system were found to be insufficient to remove or inactivate oocysts. As a result of this outbreak, the state of New York was able to amend the state pool code to include a requirement for supplemental disinfection (e.g., UV, ozone) on state spray parks; the first such state in the United States to do so. These outbreaks underscore the need for regulatory entities to review pool codes and be vigilant during plan reviews to ensure that existing regulations cover all public recreational water pools, fountains, and other water-based attractions that, regardless of design intent, could expose users to contaminated water. Lessons learned from treated venue-associated outbreaks include: 1. The potential for large-scale transmission is high. 2. The simple fecal-oral transmission mode and chlorine resistance of Cryptosporidium increase the likelihood of outbreaks becoming communitywide. 3. Combined filtration systems can lead to cross contamination of all cofiltered pools following fecal contamination. 4. Closing pools during outbreaks will lead to swimmers going to other pools, which is likely to expand the outbreak. 5. Hyperchlorination for Cryptosporidium is now the standard remediation method for disinfected aquatics venues. 6. Employees ill with AGI will continue to swim if leave or diarrhea exclusion policies are not in place. 7. Use of municipal water to fill “kiddie” pools without further disinfection or filtration (i.e., essentially untreated water) has been associated with disease transmission. 8. Satellite swimming-associated outbreaks can occur after other outbreaks of enteric illness if the population is not well educated about the risks of swimming when ill with diarrhea. 9. Swimming while traveling internationally can be associated with disease transmission 10. Interactive fountains may not be designed to adequately treat water. 11. Interactive fountains are not covered by all pool codes.
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Recreational Water Types and Prevention
It is clear from the breadth and number of outbreaks reported that Cryptosporidium has become a major international public health challenge, particularly in treated swimming venues. Adoption of a uniform and risk-based approach to reducing transmission is greatly needed (Dadswell, 1996; Pool Water Treatment Advisory Group, 1999; Gregory, 2002; WHO, 2003, 2006; Nichols, 2006).
A.
Marine and Fresh Water
Data from some of the freshwater outbreaks discussed above suggest that reducing the risk of cryptosporidiosis transmission via recreational water will require a concerted, multifaceted effort involving public health, environmental protection agencies, environmental resource management, aquatics professionals, and the swimming public. In order to decrease the risk of enteric infections, freshwater-associated outbreak data supports: 1. Implementation of routine fecal indicator monitoring, particularly when more rapid testing methods are adopted. 2. Periodic completion of sanitary surveys to identify potential risks. 3. Advising swimmers to refrain from swimming after heavy rainfall events. 4. Enforcement of prohibitions on dumping of human waste from boats. 5. Monitoring use of animal waste on watershed land near bathing beaches with enforced prohibition when warranted. 6. Prohibiting use of untreated mineral water in artificial, full-body recreation facilities without continuous filtration and disinfection. In natural environments, reduction of oocyst concentrations through improved watershed management and reductions in oocyst introduction from point and nonpoint sources of animal and human contamination is necessary. EPA criteria for microbial fecal indicator testing of natural recreational waters was published in 1986 (EPA, 1986) and promulgated in 2006. Although the percentage of beach closure days is small relative to the total (4%), the continued contamination of public beaches in the United States suggests that fecal indicator testing, in some form, will continue to be necessary in the future. EPA development and validation of a new generation of fecal indicator tests that can be performed in hours instead of days would allow beach managers to directly protect the health of beachgoers on the same day that contamination is detected (Wade et al., 2006). Such rapid tests should also decrease the number of unnecessary closures which occur because the test takes 24 to 48 h to complete. Decreasing external contamination of recreational water by sewage, watershed run-off, etc., is a complex problem that will require continued vigilance by EPA, public health, and environmental health partners. WHO has released guidelines for safe coastal and freshwater beaches that rely on water quality monitoring and development of a Code of Good Practice (WHO, 2003). In addition to external contamination of natural recreational beaches, beach and resource managers must be cognizant of possibilities for contamination within their own beach areas. Sanitary surveys to identify modes of potential contamination such as leaking septic tanks, inappropriate dumping of recreational camper waste, etc., will be needed in order to protect future water quality. Natural swim areas that have poor circulation (e.g., shallow, little wave action) may need to develop engineered solutions to improve the amount of water circulation (e.g., pumping of water across swim areas) in order to circulate and dilute more focal bather contamination that is unlikely to be detected by routine fecal indicator testing. Beaches may also consider other physical barriers to potential microbial contamination. Use of untreated hot spring water in full-body family swimming recreation increases the risk for enteric disease transmission, including cryptosporidiosis, and the practice should be discouraged.
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Cryptosporidium has become a major international public health challenge in treated swimming venues. Although it is likely that most potential outbreaks of enteric illness are prevented by appropriate disinfection and pH control, the emergence of Cryptosporidium is creating a paradigm shift in thinking about pool disinfection. The reliance on two major treatment barriers, filtration and halogen disinfection, over the past 80 years is no longer sufficient to adequately protect public health. Improved training, maintenance, and pool operation will continue to be critical to preventing future outbreaks, but it will not be the complete answer to preventing Cryptosporidium transmission (Dadswell, 1996; Pool Water Treatment Advisory Group, 1999; Gregory, 2002; Kebabjian, 2003; WHO, 2006; Kirkpatrick et al., 2006; Nichols, 2006). Recommendations for treated venue-associated outbreaks include: 1. Filtration systems, particularly those for “kiddie” pools, should be kept separate to reduce cross-contamination of other cofiltered pools. 2. Pool remediation (e.g., hyperchlorination) should be conducted, and the pool rapidly reopened for public use to minimize contamination of other facilities. 3. Sick leave or alternative duty policies for aquatics staff ill with AGI should be developed. 4. The use of temporary, untreated pools (inflatable pools, paddling pools) filled with municipal water should be discouraged and should be prohibited in institutional settings (e.g., day care centers). 5. Extensive education should be undertaken during and after enteric illness outbreaks to inform the public of the potential for satellite outbreaks to occur if people swim when ill with diarrhea. 6. International travelers should be informed about the potential for disease transmission associated with recreational water. 7. All new water feature designs should be reviewed to ensure that adequate water treatment is included. 8. Pool codes should be periodically reviewed and updated to ensure coverage of new water feature designs. It is likely that a combination of changes in pool water treatment will be required to reduce the transmission of enteric pathogens, including Cryptosporidium. Improving existing technology is important but inadequate to solve the challenge. Improving filtration through use of alternative media (e.g., diatomaceous earth, cellulose) or technologies (membrane filtration) that are more effective at Cryptosporidium removal will be advantageous (Nieminski and Ongerth, 1995; Ongerth and Hutton, 1997, 2001; Betancourt and Rose, 2004). However, this may not be totally preventive because even pools with diatomaceous earth filters can still be associated with outbreaks (Sorvillo et. al., 1992; Beach et al., 1996; Causer et al., 2006). Boosting the efficiency of filtration through operational use of coagulants or flocculants in pool water has been demonstrated to improve the efficiency of filtration for Cryptosporidium and can be applied to swimming pools (States, 2002; Betancourt and Rose, 2004). The addition of in-line supplemental disinfection with established technologies that inactivate Cryptosporidium, such as ozone (Korich et al., 1990; Corona-Vasquez et al., 2002b; Betancourt and Rose, 2004) or UV irradiation (Bukhari et al., 1999; Clancy et al., 2000, 2004; Craik et al., 2001; Betancourt and Rose, 2004; Rochelle et al., 2005) complements existing halogen disinfection and filtration and is likely to become an accepted third level of treatment in the pool industry. Systems for use in the aquatics industry exist for both these technologies. However, these technologies are only supplemental to existing halogen disinfection because neither leaves a critical level of residual disinfectant in the water after treatment. The systems are installed to feed chemical into, or irradiate, the water stream as it leaves the filter to return to the pool. As a result, disinfection by these systems is dependent on the turnover rate of the pool (the length of time for an equivalent pool volume of water to pass through the filter system), which can typically range from 30 min to 6 h, depending on pool volume. Therefore, a continuously recirculating pool theoretically needs at least four to six pool volume turnovers to treat all the water in the pool. This dependence on pool circulation rates suggests that these technologies should reduce the long-term buildup of parasite in the
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pool water and filter, and should decrease the length of time for transmission of the parasite following fecal contamination. However, these systems are unlikely to fully prevent transmission because of the time needed for complete pool turnover (typically 2 to 24 h); in fact, some outbreak-associated pools have had ozone systems installed (Hunt et al., 1994; PHLS, 1999c; Louie et al., 2004). One outbreak documented that the side-stream ozonation system was known to be malfunctioning (PHLS, 1999c); it is unknown whether the other systems were performing to design specifications. Newer technologies such as chlorine dioxide, which inactivates Cryptosporidium (Corona-Vasquez et al., 2002a) and leaves a residual disinfectant level in the water, need to be further tested for their long-term feasibility and safety in the pool environment. Other pool design changes should include separate filtration systems for high risk bather areas (i.e., kiddie pools) to avoid cross-contamination of bathing areas. In addition, it is important that facilities include optimal numbers, placement, and design of sanitary facilities (e.g., restrooms, showers, diaper-changing areas) to encourage usage. Pool operation practices that could impact Cryptosporidium transmission and should be considered include: 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11.
Uniform, risk-based pool operation codes or regulations Uniform, risk-based inspection criteria Regular pool water replacement that is dependent on bather load Regular shocking of the pool water to inactivate pathogens and oxidize organic material Mandatory training for pool operators Employee illness policies that place ill workers in non-water-related activities Regular equipment maintenance, system operation, and water quality checks Disinfection guidelines for body waste and body fluids disinfection Clear communication channels for employees to report or resolve operation problems Mandatory sanitary showers before pool entry Regular swim breaks, particularly in children’s areas, to encourage restroom use and appropriate diaper changing 12. Establishment of effective communication networks with other aquatics venues, day care centers, and health departments so that increases in disease case reports can be readily and quickly communicated, allowing preventive actions to be implemented on a communitywide basis. Based on the experience gained from outbreaks associated with disinfected venues, it seems prudent that in the event of a potential cryptosporidiosis outbreak (e.g., increased case reporting over baseline, increased ill customer complaints) pools should be preemptively closed, hyperchlorinated (CDC, 2001c), water quality brought back within regulatory guidelines, and rapidly reopened for public use. Closure could include implicated pools or all community pools depending on the situation and potential for multipool transmission. Such outbreaks have the potential for amplification into communitywide outbreaks. To reduce this risk, a communication system could be established that would allow public health officials to contact key institutions (e.g., pools, day care centers, health institutions) when increases in enteric illness are detected at the community level. Proactive, prevention measures could then be implemented. Raising awareness of swimmers about the potential for disease transmission could decrease the documented numbers of ill persons who choose to continue swimming while ill with diarrhea. Regardless of design changes or operation and management improvements, contamination will continue to occur without a concerted effort to inform and educate the public about the infectious disease issues associated with communal swimming. The ease with which fecal contamination of pools can occur, combined with the high bather loads and the presence of incontinent swimmers (e.g., diaper-aged children), increases the risk for spreading infectious disease through recreational water use. This is a strong incentive for raising awareness about recreational water illnesses within the general swimming public. Development of effective health communication materials for the public is essential to getting
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swimmers to understand that swimming in a communal bathing environment can transmit a variety of pathogens. Strong messages coming from public health officials and health-care practitioners (e.g., pediatricians) are essential to reduce the continued use of swimming venues by persons ill with diarrhea. Because of the known postdiarrhea excretion of oocysts (Chappell et al., 1996), variation exists as to whether messages about refraining while ill with diarrhea should also include some period of exclusion after resolution of symptoms. No data exists on whether outbreaks are caused by diarrheal events or formed stool events (extended oocyst excretion), although cryptosporidiosis diarrhea has a higher titer of oocysts than formed stool from infected persons (Chappell et al., 1996). However, diarrheal events are suspected, based on the high attack rates observed. In addition, formed fecal accidents from swimming pools were not found to contain Cryptosporidium (Giardia was found in 4.4%) in one study (CDC, 2001b). Formed stools are also more likely to be observed and removed before dispersion in a pool and would therefore be expected to have a reduced risk of causing outbreaks (CDC, 2001b, 2001c; WHO, 2006). Based on this rationale, practicality would suggest that general messages for all swimmers would focus on diarrhea as the main risk factor and key to preventing large-scale transmission. General messages about not swimming when ill with diarrhea, reducing the swallowing of potentially contaminated water, and practicing good hygiene before and during swimming activities will be essential for reducing transmission. Because of the longer excretion times for some waterborne pathogens, more specific messages should be used for those persons diagnosed with these illnesses, including cryptosporidiosis (e.g., do not swim while ill with diarrhea nor for 2 weeks after resolution of symptoms). These messages can be part of physician anticipatory guidance to ill patients (American Academy of Pediatrics, 2006) or during outbreaks of known waterborne enteric disease such as cryptosporidiosis. Additionally, persons with a weakened immune response should be warned about the potential for transmission of cryptosporidiosis through recreational water use (Kaplan et al., 2002).
VI. Perspective The public challenge created by Cryptosporidium’s biology, environmental hardiness, and halogen resistance has been well documented. As a result, the parasite has become the major cause of AGI associated with disinfected swimming venues in developed countries. Since 1988, eight countries have documented 136 outbreaks of cryptosporidiosis (91.2% [124] in disinfected water, 8.8% [12] in freshwater) that have sickened at least 19,271 people (95.8% [18,472] in disinfected water, and 4.2% [799] in freshwater). The potential for large-scale transmission is highlighted by having a single outbreak that sickened over 5000 people; 25 outbreaks each infected more than 100 people and four outbreaks each infected over 1000 people. Multiple outbreaks linked to swimming have expanded into communitywide outbreaks. Current accepted pool disinfection systems are generally inadequate to inactivate Cryptosporidium. Pool operation and operator training is deficient in many cases and increases the risk of infectious disease transmission. Existing swimming behaviors (e.g., swallowing water, not refraining from swimming while ill with diarrhea) violate basic hygiene and disease control maxims. The complexity of the problem makes it important that a multifaceted approach to prevention be developed. Such an approach will involve: 1. 2. 3. 4. 5. 6. 7.
Updated, risk-based regulation and enforcement practices at public swimming areas Improved design of public swimming areas Expanded training of aquatics staff Upgraded disinfection and filtration systems Improved operation, monitoring, and maintenance standards Optimized operational policies and management practices Expanded disease communication networks that include pools and other vulnerable institutions
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8. Communitywide education of swimmers about basic hygiene practices and disease control behaviors needed at public swimming areas. Implementation of such a prevention program will need a commitment of resources and staff on a sustained basis to reverse current trends and reduce the risk of future disease transmission.
Acknowledgments I would like to thank Jonathan Yoder for assistance with the WBDO database and figure and table development. I would also like to thank my other WBDOSS colleagues, Sharon Roy, Rebecca Calderon, and Gunther Craun, for past and present interaction with the surveillance system.
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Smith-Palmer, A. and Cowden, J. 2003. Annual report of general outbreaks of infectious intestinal disease in Scotland, 2002. SCIEH Wkly. Rep. 37(22), 141–143. Smith-Palmer, A. and Cowden, J. 2004. Annual report of general outbreaks of infectious intestinal disease in Scotland, 2003. SCIEH Wkly. Rep. 38(32), 190–194. Smith-Palmer, A., Cowden, J. and Locking, M. 2005. Annual report of general outbreaks of infectious intestinal disease in Scotland, 2004. HPS Wkly. Rep. 39(50), 282–285. Sopwith, W., Osborn, K., Chalmers, R. and Regan, M. 2005. The changing epidemiology of cryptosporidiosis in North West England. Epidemiol. Infect. 133, 785–793. Sorvillo, F.J., Fujioka, K., Nahlen, B., Tormey, M.P., Kebabjian, R. and Mascola, L. 1992. Swimmingassociated cryptosporidiosis. Am. J. Pub. Health. 82, 742–744. Stafford, R., Neville, G., Towner, C. and McCall, B. 2000. A community outbreak of Cryptosporidium infection associated with a swimming pool complex. Commun. Dis. Intell. 24, 236–239. States, S.M., Tomko, R., Scheuring, M. and Casson, L., 2002. Enhanced coagulation and removal of Cryptosporidium. J. AWWA 94, 67–77. Stehr-Green, J.K., McCaig, L., Remsen, H.M., Rains, C.S., Fox, M. and Juranek, D.D. 1987. Shedding of oocysts in immunocompetent individuals infected with Cryptosporidium. Am. J. Trop. Med. Hyg. 36, 338–342. Stevenson, A.H. 1953. Studies of bathing water quality and health. Am. J. Pub. Health. 43, 529–538. Sundkvist, T., Dryden, M., Gabb, R., Soltanpoor, N., Casemore, D. and Stuart J. 1997. Outbreak of cryptosporidiosis associated with a swimming pool in Andover. Commun. Dis. Rep. CDR Rev. 7(12), R190–192. Tamburrini, A. and Pozio, E. 1999. Long-term survival of Cryptosporidium parvum oocysts in seawater and in experimentally infected mussels (Mytilus galloprovincialis). Int. J. Parasitol. 29, 711–715. Tate, D., Mawer, S. and Newton, A. 2003. Outbreak of Pseudomonas aeruginosa folliculitis associated with a swimming pool inflatable. Epidemiol. Infect. 130, 187–192. Tillett, H.E., de Louvois, J. and Wall, P.G. 1998. Surveillance of outbreaks of waterborne infectious disease, categorizing levels of evidence. Epidemiol. Infect. 120, 37–42. Uga, S., Matsuo, J., Kono, E., Kimura, K., Inoue, M., Rai, S.K. and Ono K. 2000. Prevalence of Cryptosporidium parvum infection and pattern of oocyst shedding in calves in Japan. Vet. Parasitol. 94, 27–32. U.S. Bureau of the Census. Statistical Abstract of the United States: 1995. 115th ed. Washington, DC, US Bureau of the Census, 1995. van Asperen, I.A., de Rover, C.M., Schijven, J.F., Oetomo, S.B., Schellekens, J.F., van Leeuwen, N.J., Colle, C., Havelaar, A.H., Kromhout, D. and Sprenger, M.W. 1995. Risk of otitis externa after swimming in recreational fresh water lakes containing Pseudomonas aeruginosa. BMJ 311, 1407–1410. Visvesvara, G.S. and Stehr-Green, J.K. 1990. Epidemiology of free-living ameba infections. J. Protozool. 37, 25S–33S. Wade, T.J., Calderon, R.L., Sams, E., Beach, M.J., Brenner, K.P., Williams, A.H. and Dufour, A.P. 2006. Rapidly measured indicators of recreational water quality are predictive of swimming-associated gastrointestinal illness. Environ. Health Perspect. 114, 24–28. Ward, P.I., Deplazes, P., Regli, W., Rinder, H. and Mathis, A. 2002. Detection of eight Cryptosporidium genotypes in surface and waste waters in Europe. Parasitology 124, 359–368. Wheeler, C., Vugia, D., Thomas, G., Beach, M.J., Carnes, S., Maier, T., Gorman, J., Xiao, L., Arrowood, M., Gilliss, D. and Werner, S.B. 2007. Outbreak of cryptosporidiosis at a California waterpark: employee and patron roles and the long road towards prevention. Epidemiol. Infect. 135, 302–310. Wiedenmann, A., Kruger, P., Dietz, K., Lopez-Pila, J.M., Szewzyk, R. and Botzenhart, K. 2006. A randomized controlled trial assessing infectious disease risks from bathing in fresh recreational waters in relation to the concentration of Escherichia coli, intestinal enterococci, Clostridium perfringens, and somatic coliphages. Environ. Health Perspect. 114, 228–236. Wilberschied, L. 1995. A swimming-pool-associated outbreak of cryptosporidiosis. Kans. Med. 96, 67–68. World Health Organization. 2003. Guidelines for safe recreational water environments. Volume 1, Coastal and fresh-water. Geneva, Switzerland, World Health Organization. World Health Organization. 2006. Guidelines for safe recreational water environments. Volume 2, Swimming pools and similar environments. Geneva, Switzerland, World Health Organization. World Waterpark Association. 2006. Waterpark industry general and fun facts. http://www.waterparks.org/other Articles/Generalfacts.pdf (accessed January 6, 2007).
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Xiao, L., Fayer, R., Ryan, U. and Upton, S.J. 2004. Cryptosporidium taxonomy: recent advances and implications for public health. Clin. Microbiol. Rev. 17, 72–97. Xiao, L., Singh, A., Limor, J., Graczyk, T.K., Gradus, S. and Lal, A. 2001. Molecular characterization of cryptosporidium oocysts in samples of raw surface water and wastewater. Appl. Environ. Microbiol. 67, 1097–1101. Yoder, J., Blackburn, B., Levy, D.A., Craun, G.F., Calderon, R.L. and Beach, M.J. 2004. Surveillance for recreational water-associated outbreaks—United States, 2001–2002. MMWR Surveill. Summ. 53(No. SS8), 1–21. Yokoi, H., Tsuruta, M., Tanaka, T., Tsutake, M., Akiba, Y., Kimura, T., Tokita, Y., Akimoto, T., Mitsui, Y., Ogasawara, Y. and Ikegami, H. 2005. Cryptosporidium outbreak in a sports center. Jpn. J. Infect. Dis. 58, 331–332.
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13 Waste Management
Dwight D. Bowman
CONTENTS I. II.
Introduction .................................................................................................................................. 371 Wastewater Effluent and Its Treatment ........................................................................................ 372 A. Effluent Treatment: General............................................................................................ 373 B. Effluent Treatment: Ozonation........................................................................................ 374 C. Effluent Treatment: UV Light......................................................................................... 374 D. Effluent Treatment: Chlorine Dioxide ............................................................................ 374 E. Effluent Treatment: Sand Filtration ................................................................................ 375 III. Removal by Clarifiers .................................................................................................................. 375 IV. Biological Treatment of Wastewater............................................................................................ 376 V. Overall Effects of Wastewater Treatment .................................................................................... 376 VI. Sludge Treatment.......................................................................................................................... 376 A. Anaerobic Digestion Treatment ...................................................................................... 378 B. Aerobic Digestion Treatment.......................................................................................... 378 C. Composting Treatment .................................................................................................... 378 VII. Dairy Cattle Manure and Treatment ............................................................................................ 378 A. Inactivation within Calf Facilities................................................................................... 379 B. Anaerobic Digestion Treatment ...................................................................................... 379 C. Biodrying and Compost Treatment................................................................................. 380 D. Oocysts in Manure Applied to Soil ................................................................................ 380 VIII. Beef Cattle Manure and Treatment.............................................................................................. 381 A. Composting Treatment .................................................................................................... 381 B. Runoff Control ................................................................................................................ 381 IX. Swine Manure and Treatment ...................................................................................................... 381 A. Lagoon Treatment ........................................................................................................... 382 B. Anaerobic Digestion Treatment ...................................................................................... 382 X. Summary and Conclusions........................................................................................................... 382 References........................................................................................................................... 383
I.
Introduction
Manure and sewage management requires that both the solid and liquid phases of waste be treated or processed. In waste treatment systems, there may or may not be partitioning of the waste into separate liquid and solid waste streams. In the treatment of human sewage, there is almost always a partitioning into a sludge stream (the solids) and a liquid stream (the effluent); this is due mainly to the collection system, which is based on using wastewater to carry the solid portions of the sewage to treatment centers. In the case of cattle, manure is often collected and processed as a single relatively dry solid matrix. Pig manure is often handled in slurries and may or may not be partitioned for treatment into solid and
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effluent streams. Even when relatively dry cattle manure is processed, there is often a dewatering phase that will produce a liquid effluent. Oocysts of Cryptosporidium can enter either phase of the partitioned waste stream. If a great deal of liquid is removed in order to disinfect the solids, the majority of the oocysts might be transferred into the liquid stream and released untreated. Thus, processing of waste must always consider the treatment of both the solids and liquid. The goals of waste management in the human sewage industry often differ from management of manure on farms. A major goal of waste treatment and management with sewage is to clean and disinfect the liquid phase so that clear, pathogen-free water can be returned to the environment. The solids portion of sewage is often treated with processes that will reduce the number of pathogens or destroy them. Solids are termed sludge before processing and biosolids after processing. They may then be land-applied or disposed of in landfills. The goal of treating animal manure often is to reduce pathogens before application to crop land. There was a time when manure was simply hauled from barns to fields for use as fertilizer and soil emendation. Now, manure is often treated to reduce the number of pathogens, to prevent runoff into surrounding surface waters and percolation into ground waters. This chapter will discuss the treatment methods for the inactivation and removal of oocysts in different process streams. Oocysts can be present in effluents, solids, and manures (cattle and pigs). Of course, with range cattle and sheep on pastures, treatment becomes a matter of runoff control rather than the processing of waste from animals held in concentrated animal feeding operations (CAFOs). For pastures, much pathogen control is through best management practices that focus on preventing excessive pasture runoff and developing riparian barriers that restrict the flow of pathogens or slow their movement from fields into adjacent water courses. This latter aspect of control has dealt mainly with the prevention of nutrient runoff and has generated a giant body of literature and research directed primarily by the USDA’s Natural Resources Conservation Service (originally called the Soil Conservation Service). For discussion of runoff prevention and the different best management practices, readers are referred to their Web site (http//www.nrcs.usda.gov).
II.
Wastewater Effluent and Its Treatment
Sewage treatment plants can produce three or more types of effluents, usually named primary, secondary, and tertiary effluents. Primary effluents are those specifically recovered from clarifiers, in which there is little water treatment other than the settling of solids to separate the water portion. In most modern plants there is some form of secondary treatment applied to primary effluents before release into the environment. (Some plants have no primary clarification. All water entering these plants goes into some form of treatment immediately upon entering the plant; therefore, these plants have no primary effluent.). Secondary wastewater treatments consist of processes such as activated sludge treatment, in which water with low solids content is mixed for some period before settling; trickling filters, in which the water is passed over large tanks containing biofilms on various support media before settling; and extended aeration, in which water and solids are mixed with air for extended periods before settling. Water treated with secondary processes might then be treated with various tertiary processes, not all of which are for pathogen removal or disinfection. Tertiary processes consist of treatments such as phosphorus removal, denitrification, sand filtration, ozonation, or UV radiation. Sometimes, plants utilize more than one of these tertiary treatments before releasing the water into the environment. Some plants chlorinate the final effluent, before or after other tertiary processes. Often, there is a dechlorination step before the finished water is released into the waterway. Thus, it is very difficult to compare effluents among plants, because for various design reasons they may markedly differ in what happens to the wastewater from the time it enters the treatment plant until it leaves. Some processes such as phosphorus removal involve a flocculation process and settling that will also likely reduce particulates (including oocysts) that leave the plant in the effluent stream. Also, sand filters can cause marked reductions in oocyst numbers leaving a plant. Disinfection processes such as UV light treatment and ozonation can kill oocysts although they remain intact, and nonviable oocysts will then enter the effluent stream, where they can then be detected by routine recovery methods. Thus, it is possible that plants releasing killed intact oocysts in the final
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effluent can appear to be releasing more oocysts than a plant that has no disinfection process but which releases a few oocysts. Until a method is developed that is capable of rendering 100% of the oocysts noninfectious, or a real-time detection method is developed to distinguish infectious oocysts from noninfectious ones, it will be very hard to define what the recovery of oocysts in effluent samples actually means relative to environmental release of infectious organisms. Despite treatment with the aforementioned processes, oocysts are omnipresent in sewage treatment plant effluents. Sampling is problematic because oocysts are relatively small and difficult to recover, and it is not possible to determine the species of each recovered oocyst. Examination of secondary effluents (following activated sludge treatment) from seven treatment plants in Arizona revealed oocysts in all the effluents; the numbers ranged from 140 to 3960 oocysts per liter (Madore et al., 1987). Two plants that had sand filters following the activated sludge treatment had the lowest numbers of oocysts. In all of four treatment plant effluents in Israel, the oocyst numbers ranged from 8.2 to 270 per liter (Zuckerman et al., 1997). Oocysts were found in seven treated sewage effluents in England (Bukhari et al., 1997) and in one of four sewage effluents from Scotland (Smith et al., 1991). In Turin, Italy, sampling of the Azienda Po-Sangone sewage treatment works (having both primary and secondary treatment, with tertiary treatment consisting of dephosphorylation and multilayer (gravel, sand, and carbon) filtration, all 11 effluent samples contained oocysts, with a mean of 0.2 oocysts per liter (Carraro et al., 2000). In Rome, 90% of repeated samples from a secondary clarifier following an activated sludge treatment tank contained oocysts in the effluent, with numbers ranging from 0 to 82 oocysts per liter (Bonadonna et al., 2002). In France, effluents from nine treatment plants with activated sludge systems contained from 0.4 to 209.2 oocysts per liter (Lemarchand and Lebaron, 2003). In five treatment plants in Spain, all secondary effluents contained oocysts, whereas only three-fourths of the samples of tertiary effluents contained oocysts (Montemayor et al., 2005). Of 29 effluent samples from 12 sewage treatment plants in the Sydney, Australia, drinking water catchment, 76% contained oocysts (a median of 0.7 per liter), with a maximum recovery of 290 per liter (Charles et al., 2003). Following an outbreak of human cryptosporidiosis in Sydney, only one of three effluent samples from the Manley Sewage Treatment Plant was found to have oocysts (3 per 10 L), whereas two of six influent samples contained oocysts (Wohlsen et al., 2006). In Norway, samples of effluent were collected from three sewage treatment plants; 72 samples were collected over a number of months, and 61% were positive for oocysts with the number of oocysts, ranging from 100 to 44,500 per liter (Robertson et al., 2006). Only a few studies have reported genotyping of oocysts in effluent. Oocysts from a plant in Switzerland were identified as C. muris; those from one German plant were identified as C. parvum, and those from two other German plants were identified as C. hominis (Ward et al., 2002). In Japan, oocysts from sewage treatment plants (whether from influent or effluent samples is not stated) were found to be C. parvum, C. hominis, C. suis, and Cryptosporidium mouse genotype (Hashimoto et al., 2006). In upstate New York, effluents entering the Cayuga Lake watershed contained five genotypes of Cryptosporidium in water from six of seven wastewater treatment plants sampled (Lucio-Forster, 2006). The isolates identified grouped with three Cryptosporidium species, C. andersoni, C. baileyi, and C. parvum, and with two storm water genotypes, W4 (a cervine genotype found frequently in deer, preweaned lambs, and postweaned calves, and occasionally in humans) and W12 (a wildlife genotype).
A.
Effluent Treatment: General
Effluents from wastewater treatment systems can be treated to destroy oocysts. Effluent streams are often quite clear of visible particulates, and thus amenable to treatment with processes similar to those used to treat drinking water. Resistance of Cryptosporidium oocysts to chlorine levels used in drinking water is one reason the parasite has received so much notoriety in the world of waterborne pathogens. Because utilization of slightly higher levels of chlorine at sewage treatment plants would have little or no effect on the oocysts, other tertiary disinfection treatments have been considered, tested, and put in place to increase removal or achieve disinfection.
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Ozone (O3), the reactive form of oxygen, has detrimental effects on many pathogens and has been utilized in many disinfection systems. Ozone is typically generated by passing an electric current through oxygenrich water. Ozonation has the advantages for wastewater treatment that it is a highly effective disinfectant for many organisms, produces very few disinfection by-products, can be used to treat large volumes of water, and has no lasting residual effect. Its disadvantages are that it can create bromate as a disinfection by-product, if the water contains bromide, and it has reduced efficacy in cold water (Betancourt and Rose, 2004). Ozonation of tap water at 1°C resulted in a 2 log reduction in 10 min at a concentration of 4 mg/l and in 1 min at 2 mg/L with the water at 20°C (Corona-Vasquez et al., 2002). Oocysts treated with ozone at 0.4 mg/L for 2 min at 22°C caused a 2 log reduction in only one of four replicate trials (Bukhari et al., 2000). In four additional trials there was not even a 1 log reduction in infectivity. In trials at different temperatures, it was found that infectivity was highly related to temperature, with 2-log reductions occurring with 43, 12, 8.2, 3.4, and 0.72 mg/min/L at 3, 10, 15, 20, and 30°C, respectively (Hirata et al., 2001). These values are all for finished drinking water that has much lower turbidity and biological oxygen demand (BOD) than wastewater effluents. Thus, it is expected that higher ozone concentrations would be required for effluent treatment. A recent outbreak of cryptosporidiosis associated with ozonated apple cider suggests that ozone may be less effective in samples of high turbidity or perhaps when high levels of sugars are present (Blackburn et al., 2006).
C.
Effluent Treatment: UV Light
UV light has been approved for the inactivation of oocysts in drinking water in the Long Term 2 Enhanced Surface Water Treatment Rule (LT2 rule) (www.epa.gov/safewater/disinfection/ lt2/index.html). UV light has the advantages that is does not require addition of chemicals to the water, it is highly effective in killing protozoa, it has no lasting residual effects, and it generates no identifiable disinfection by-products. The disadvantages are that it is highly susceptible to interference by flocking or turbidity (serious concerns in the case of effluents versus drinking water), and it has been difficult with current equipment to design systems that are easily monitored as to output (Betancourt and Rose, 2004). In clean water the efficacy of disinfection has been found to be anywhere from 1.5 to more than 3.2 logs of inactivation at 1 to 3 mJ/cm2 of UV light (Betancourt and Rose, 2004). As with ozone, the major concern is the negative effect of high turbidity.
D.
Effluent Treatment: Chlorine Dioxide
Chlorine dioxide is a water-soluble gas. It can be manufactured on site and has a high reactivity and oxidative potential. Chlorine dioxide has only an oxidative action with no chlorinating action to cause the formation of chlorinated hydrocarbons or other organochlorine compounds. It has a high disinfectant activity over a wide pH range, has no ammonia or ammonium compounds, provides long-term bactericidal and bacteriostatic protection, and does not form chlorinophenols. It is licensed in many countries for water treatment. Based on a series of infectivity studies (Li et al., 2001), it was shown that the contact times between oocysts and chlorine dioxide were 606 min at 1 mg/L for a 2 log reduction in viability at 1°C, 442 min at 5°C, 230 min at 13°C, and 111 min at 22°C. The highest practical chlorine dioxide dose for drinking water is about 1.1 mg/L. Thus, it was suggested that a disinfection basin with a retention time of 150 min might be able to achieve 2 logs of inactivation at 22°C with the application of 1.1-mg chlorine dioxide per liter. Of course, as with ozone and UV, the contact times may need to be greater in treated water with higher particulates or BOD. It might be possible to use higher doses of ozone in the treatment of sewage effluents, and it is also possible to treat effluents with sequential UV and ozone or UV and chlorine dioxide for maximal inactivation (Meunier et al., 2006).
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Effluent Treatment: Sand Filtration
Sand filtration is used for tertiary treatment to remove particulates from effluents after secondary treatment and some form of clarification. There is every reason to believe that filtration will be effective at some level if the functional pore size is such that it can collect the oocysts, the flow is not too extreme, and as long as the backwash used to clean the filter is not allowed by accident to mix with the finished effluent. Several studies have examined the efficiency of sand filters to remove oocysts from effluent streams. Different types of sand filters have been examined. Slow sand filters of varying depths with different flow rates were found to be excellent at oocyst removal with influents of 4000 oocysts per liter and effluents containing no detectable oocysts (Timms et al., 1995). Rapid sand filters have also been found successful in the removal of oocysts from a full-scale 900-gpm treatment plant (Nieminski and Ongerth, 1995). Sand grains of 310 µm (nominal pore size of 27 µm) were highly effective in removing the 5µm oocysts from water (Logan et al., 2001). Even grains as large as 1.4 mm in diameter (nominal pore size of 55 µm) were effective in retaining oocysts. The conclusion is that some factor other than simple filtration causes retention of the oocysts within the sand filter. Of course, the success of the process is also dependent on any coagulants that might be added before filtration (Adin, 2004). The filters are probably an excellent means of reducing oocysts in wastewater effluents. They would benefit most systems seeking to minimize pathogens and obtain the highest postfiltration efficiency for any disinfection that requires water with minimal flock or turbidity.
III. Removal by Clarifiers Clarifiers work by simple settling, with solids sinking to the bottom and liquids staying on the surface. There are some special clarifiers that work in reverse, wherein particulates associated with air bubbles are caused to float to the surface while water drains from the bottom, but these are uncommon. Thus, clarifiers function mainly based on the buoyant density of the particulates being removed. The buoyant density of the oocysts of Cryptosporidium parvum based on ultracentrifugation in Percoll solutions is around a specific gravity of 1.08 (Jenkins et al., 1997). Because oocysts are very close to the density of water, they require a considerable amount of time to settle if they are not bound up in flock or adhering to other particulates. The settling velocity of the spherical oocysts with an average diameter of around 6.6 µm that have been shown to have an average density of 1009 kg/m3 is about 0.36 µm/s (Dai and Boll, 2006). This would mean that in an hour, oocysts would move only about 1.3 mm toward the bottom of a container. Thus, as stated by Dai and Boll (2006), “it appears that free oocysts can be transported long distances unless they are trapped in low-flow environments, or they are subjected to infiltration or other barriers to flow.” Fresh and fully viable oocysts might settle almost thrice as fast, but as they age and become proportionately less viable, the average settling velocity might diminish (Young and Komisar, 2005). There is some physical removal of oocysts in clarifiers, possibly owing to settling with solids or binding to other particulates that settle in the clarifiers. The flow of oocysts through one of the largest primary clarification-only wastewater treatment plants in the world in Montreal, Canada, serving a population of about 1 million people was examined (Payment et al., 2001). During testing, this plant had a capacity of 7.6 million cubic meters per day (2 billion U.S. gallons per day). About 27% of the oocysts entering the plant were removed by clarification before the water was released form the plant into the St. Lawrence River. Oocyst counts appeared rather erratic overall compared to the other pathogens monitored, including Giardia cysts, bacteria, and viruses, and the authors felt that this was due mainly to difficulties with the methodology for detection. In a laboratory-scale activated sludge system, if oocysts were added to the primary effluent entering the system, the overall removal by primary clarification, activated sludge, and secondary clarification resulted in 98.6% removal (Stadterman et al., 1995).
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Biological Treatment of Wastewater
Most secondary treatment of the liquid stream of wastewater utilizes some form of biological treatment. Such systems include extended aeration, activated sludge treatment, and trickling filters. Common to all these systems is aeration with the production of a large number of aerobic organisms that compete with and aid in the destruction and inactivation of the anaerobic or facultatively aerobic organisms routinely found in wastewater. Besides aerobic bacteria, these systems also include numerous other organisms such as algae, diatoms, free-living protozoa (flagellates, ciliates, amoeba, testate amoebae, etc.), copepods, mites, free-living nematodes, rotifers, and many other organisms (Fox et al., 1981). Many of these organisms are autotrophs, but others are predatory. Predatory organisms, including protozoa (Acanthamoeba culbertsoni, Paramecium, Stylonychia, and Euplotes) and rotifers (Euchalanis and Brachionus), have been shown to ingest relatively high numbers of oocysts per organism when given the opportunity (Fayer et al., 2000; Stott et al., 2001, 2003). Although it has not been specifically demonstrated that ingested oocysts are destroyed or inactivated within the predators, ingestion would have the effect of removing them from the effluent in secondary clarifiers as the much larger size of the predators would allow for removal by sedimentation. It is necessary to separate the action of biological treatment from sedimentation and tertiary treatment at a given plant to examine how much of the removal is due to biological stabilization and how much is due to poststabilization clarification. In an experimental activated sludge system spiked with oocysts, reduction due solely to the activated sludge treatment was around 80%, based on infectivity to mice (Villacorta-Martinez de Maturana et al., 1992). In a pilot plant, removal of formalin-fixed oocysts in a 100L aeration tank that was followed by a 50-L clarification tank resulted in a 2-log removal of oocysts (Suwa and Suzuki, 2003). When a coagulant was added prior to the settling tank, oocyst removal increased to 3 logs.
V.
Overall Effects of Wastewater Treatment
A careful 3-year study was conducted in six sewage treatment plants in Scotland to evaluate the effects of wastewater treatment on the oocysts of Cryptosporidium (Robertson et al., 2000). One of the six plants had only primary clarification; another had primary clarification followed by trickling filters; two others had primary clarification followed by activated sludge treatment; the fifth had no primary clarification, only activated sludge treatment; and the last plant in the group had primary clarification, trickling filters, and tertiary sand filtration. Based on influent versus effluent oocyst numbers, the three primary clarifiers had a mean removal of 4 to 30% (there was great variability, with ranges from 0 to 100%). In the three plants that had secondary treatment (excluding the plants with trickling filters), the overall removal of oocysts was 28 to 98%, again with wide variability in the different samples. For all six plants, the overall removal efficiency was between 5 and 91%. The authors examined the viability of oocysts in both the influent and effluent using a vital dye assay. They also found that 33% of the incoming oocysts were viable (range 7 to 100%), and 46% of the oocysts in the effluents were viable (range 20 to 100%). They found viable oocysts in the effluents of all plants, except the one simple activated sludge plant from which few oocysts were collected. To assess the effects of the different portions of the treatment systems, the researchers placed oocysts within membrane-bound chambers in different processes of the sewage treatment plants (raw sewage, settled sewage, aerobic biological treatment, final effluent, and stored sludge) for 162 h and found no reduction in viability by the dye exclusion assay or using an in vitro excystation assay.
VI. Sludge Treatment At municipal sewage treatment plants, there are several types and levels of treatment that can be applied to sludge to produce a finished biosolid. Primary sludge tends to represent sludge collected from primary
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clarifiers, whereas secondary sludge, waste-activated sludge, or extended aeration return sludge, typically refer to sludge collected from secondary biological treatment processes. Secondary sludge tends to be mixed with the primary sludge, with the exception of any that is recirculated to the biological system from which it was derived. Sludge from tertiary sand filters is often simply returned to the head of the plant, where it is mixed with newly incoming primary influent. Sludge, primary or secondary, can be disposed of directly, land-applied or landfilled, or it can be treated by various methods, as in processes where it is mixed with lime or other materials at a high pH and held above a given temperature for some period to cause pathogen inactivation through heat and pH effects. Sludge from clarifiers can also be incinerated and disposed of as ash. Some primary sludges are treated directly by composting, usually mixed with a second organic source (e.g., leaf litter) before processing. In some situations, primary sludge is simply landfilled and used as an additional source of methane generation by landfill sites for the production of electricity. Sludges from clarifiers are also formed into biosolids through various biological processes. The two most common means are anaerobic and aerobic digestion. In anaerobic digestion, the sludge is typically held at 37°C (mesophilic digesters) in a liquid state for several weeks while it undergoes methanogenic digestion. Primary digesters might be followed by secondary digesters that might also have a long detention time. Aerobic digestion most typically occurs at ambient temperatures and looks like an activated sludge system maintained with a higher concentration of solids in the mixed liquor. As with anaerobic digestion, there may or may not be a secondary set of aerobic digesters. The biosolids produced by digesters may then be land-applied or treated by processes that further reduce the pathogens present within the material. In some situations, digesters can be maintained at higher temperatures, usually 50 to 55°C (thermophilic digestion); in these cases, the higher temperatures often inactivate many of the pathogens that might be present. Processes that further reduce pathogens include treatment with high pH and increased temperatures, pelletization, composting, heat drying, heat treatment of liquid material, beta and gamma irradiation, and pasteurization. Many of the latter processes are developed under different management structures, and all may fall under different patents. There is a multitude of such systems in place that all cause pathogen inactivation, and most of them are capable of the inactivation of oocysts when operated according to design criteria. For a process to be given national acceptance as a process to further reduce pathogens, it must be approved by the Pathogen Equivalency Committee of the USEPA. The approval of the process by the EPA means that when the process is used as approved, oocysts are likely to be destroyed or inactivated by the treatment. The advantage of treating biosolids with a process that further reduces pathogens is that they can be land-applied without any restrictions on the type of land use projected for the area of application. The product of these treatments is termed a Class A biosolid, whereas material not treated in such a fashion is considered Class B material. There are various landuse restrictions placed on material that has not undergone a process that creates a Class A product. Processed biosolids from biological stabilization processes may be land-applied as liquids or they may be dewatered. There are numerous means of removing water from processed biosolids, including vacuum filtration, centrifugation, and air drying. Often, to assist in the process, various coagulants or quantities of lime are added to the material prior to the drying process. The water that is generated from the process is typically then returned to the influent stream of the treatment plant, where it reenters the treatment stream with the raw wastewater. The dewatering may or may not inactivate contained pathogens. Inactivation by dewatering may depend on the temperatures reached, the additives mixed with the product, and the amount of time the material is processed. Some biosolids are stored for months to years after biological stabilization before land application. In some states, no land application is allowed in winter months, so stabilized biosolids are stored for months until they can be applied to the land. In some municipalities, biosolids are stored in lagoons, and some of these lagoons are large enough that the retention time of the biosolids within these lagoons can reach several years before the material is ever used for land application. In these situations, many of the pathogens are further reduced by the long-term storage, but it is very difficult to determine the actual efficacy of inactivation of oocysts in such long-term systems.
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Cryptosporidium and Cryptosporidiosis, Second Edition Anaerobic Digestion Treatment
Most oocysts that enter sewage treatment plants and partition with the solids portion of the process will undergo anaerobic digestion if the primary sludge is so treated. Mesophilic anaerobic digestion caused an 80% reduction in the viability of oocysts in 3 days, but a few were still viable after 18 days (Whitmore and Robertson, 1995). Mesophilic digestion also has been reported to have caused a 90% inactivation within 4 h and a 99.9% inactivation within 24 h (Stadterman et al., 1995). A 99% inactivation (2-log inactivation) of oocysts did not occur until 10 days after the oocysts were added to mesophilic anaerobic digesters (Kato et al., 2003); at 47°C and 55°C, 2 logs of inactivation were observed at 4 days and 2 days, respectively. It appears that there is no reduction in the number of detectable oocysts when sludge is treated by anaerobic digestion with typical detention times of several weeks (Chauret et al., 1999). It is to be expected that oocysts held for any time at 55°C or at higher temperatures will be inactivated.
B.
Aerobic Digestion Treatment
Aerobic sludge digestion is used in a number of municipal sewage treatment plants, although they are usually smaller systems. The inactivation of oocysts in bench-top aerobic digesters was examined by Kato et al. (2003). They found that at 37, 47, and 55°C, the oocysts were inactivated in a fashion very similar to oocysts in anaerobic systems, with 2 logs inactivation occurring at 8 days at 37°C, 4 days at 47°C, and very soon after they were added at 55°C. Inactivation was monitored by dye exclusion.
C.
Composting Treatment
Composting is the aerobic stabilization of solid material. Biosolids treatment requires that specific temperatures be achieved and that compost piles be turned at appropriate intervals to ensure that all sections of the pile achieve maximal temperature. Again, because compost typically reaches temperatures above those where most proteins are usually denatured, there is reason to believe that the infectious sporozoites contained in the oocysts would be killed at these temperatures. The concern is usually one of uniformity of treatment. One study found significant numbers of oocysts present even after 30 weeks of composting, with 10% of the samples containing oocysts (Rimhanen-Finne et al., 2004). In fact, the number of oocysts found after 10 weeks of composting was the same as after 30 weeks of composting. However, no attempt was made to assess the effects of the process on the viability of the oocysts, and there is reason to believe that they were probably inactivated by the temperatures in the composting process if the compost pile was properly mixed and its temperature monitored appropriately.
VII. Dairy Cattle Manure and Treatment Dairy cattle manure is handled differently from that of beef cattle. Most beef cattle spend their early lives on pastures with their dams. Beef cattle only come together in large numbers for brief periods in feedlots, where massive amounts of feces accumulate in relatively short periods. In the larger modern dairy farms, cattle rarely see pasture, either as calves or adults. Throughout the life of a dairy cow, it produces manure that must be collected and disposed of in some fashion. In some systems it is collected by water, using different wash systems. In other methods, it is kept relatively dry as it is moved from one place to another on a farm. The processing of cattle manure to reduce pathogens is to some extent a fairly new phenomenon, at least in terms of scale. Large volumes of animal manure have been handled and processed in different ways for many years for sanitation. It was only about 100 years ago that the only means of transportation other than steam-powered trains and ships were horse-drawn conveyances. Cities used to contend with literally hills of horse manure well into the 1920s. This manure had to be hauled to a disposal site, where it was treated, probably with lime, to minimize its attractiveness to flies. Today, large concentrations of horses in any area around the world are usually relatively rare occurrences. The opposite has occurred in the world of dairy farming. Until recently, most dairy farmers had only 50 or 60 milking cows. Farms
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now can have more than 5000 milking cows. Previously, farms tended to have crops planted for feeding cattle and for human consumption. Manure was considered a form of fertilizer. Now, with large-scale farms, much feed is grown off site, and the farm may not be directly supporting any crop production. Thus, the manure requires disposal. The public has also become very concerned about potential manurerelated illness. Therefore, farms are finding it necessary to develop means of treating the manure before it is disposed of or land-applied. This has led to a recent increase in means of processing waste from cattle before its disposition or land application. Dairy waste is markedly different from human waste in how it is collected. A relatively large quantity of water is used to collect human excreta from households in municipal collection systems, whereas dairy waste is often collected without addition of water, and even when water is added, the solids contents remains much higher than is ever the case with sewage treatment plant influents. Methods to manage dairy waste include composting, anaerobic digestion (also applicable to methane generation for energy recovery), anaerobic lagoons, aerobic ponds, and artificial wetlands. Furthermore, a major discovery in Cryptosporidium taxonomy associated with dairy manure is the discovery that C. parvum, the primary zoonotic species, is excreted primarily by young (preweaned) calves, whereas older dairy cattle shed oocysts predominantly of the nonzoonotic species, C. bovis, C. andersoni, and the Cryptosporidium deer-like genotype (Fayer et al., 2005, 2006). Thus, the zoonotic threat relative to cryptosporidiosis from dairy waste is primarily associated with manure from very young calves. However, because of potential bacterial pathogens associated with dairy manure, treatment of all dairy waste is necessary before it can be released into the environment. Processes that induce several logs of inactivation of oocysts are likely to also serve to protect the public from these other pathogens.
A.
Inactivation within Calf Facilities
For health reasons, many young dairy calves are housed in ways that provide shelter from the environment, separation of animals, good drainage, adequate bedding, and ventilation without drafts. Some housing methods include outdoor hutches, greenhouses, pens, and tie stalls. The goal is to minimize the spread of infectious agents between calves, from older cattle to young calves, and to protect the general environment from contamination with pathogens from sick calves. In some of these systems, the calves are often housed for several weeks in the same area with the regular addition of fresh bedding. Typically, the pack that builds up under a calf is not removed until the calf grows to an age where it can be transferred to group housing. The manure from the calves is then, very often, mixed with and disposed with manure from adult cattle. Based on the recent taxonomic recognition that C. parvum primarily infects and is excreted by preweaned calves, it would be more appropriate to treat manure from young calves separately. Inactivation of oocysts of C. parvum in conventional tie stalls and solar calf housing systems was examined by Collick et al. (2006). Oocysts were placed in chambers mounted in protective disks on the floors under the calves in the two types of housing facilities. Viability of the oocysts, examined at 4, 6, and 8 weeks, was reduced from 15 to 30% in the winter months to less than 10% in the warmer months. No difference was found in the rate of oocyst inactivation in the two systems, but in the solar housing system it was relatively easy to keep the calf manure isolated from the other manure for additional treatment and pathogen inactivation. Survival of oocysts in manure piles held in fall and winter conditions was examined by Jenkins et al. (1999). These piles were simple storage piles, not turned or aerated as they would be if they were maintained as compost piles. However, even without turning, the internal temperatures of the piles reached between 30 and 32°C and a high percentage of oocysts (80 to 95%) were inactivated. It would be expected that higher rates of inactivation would occur if the piles were stored under similar conditions during summer months.
B.
Anaerobic Digestion Treatment
Inactivation of oocysts in cattle manure undergoing mesophilic (38°C) and thermophilic (55°C) anaerobic digestion in a series of digesters was examined by Garcés et al. (2006). Mesophilic digestion reduced
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oocyst infectivity by 2 logs after 4 h. Loss of infectivity was total after 4 h in the 55°C digester, 6- or 7-log inactivation based on the detection limit of the cell culture assay used for the infectivity. Inactivation was also complete when the oocysts were treated with the two digestion processes used in series. The effect on oocysts in anaerobic facultative lagoons containing dairy wastewater (after removal of the solids fraction by physical separation) was examined by Karpiscak et al. (1999, 2001). There was an increase in the number of oocysts present in the liquid from the dewatered wastewater, but there were no detectable oocysts present after treatment of the liquid in anaerobic facultative lagoons (200 m × 33 m × 5 m that normally operated at depths of 4 to 4.5 m). Based on the incoming loading of 2.64 × 103 oocysts per 100 mL, it was calculated that the removal rate was greater than 3 logs.
C.
Biodrying and Compost Treatment
Biodrying is a form of composting that uses forced aeration along with the heat generated by natural aerobic decomposition to dry the manure mix. Collick et al. (submitted) examined a biodrying process for the treatment of dairy cow manure using eggs of the pig roundworm, Ascaris suum, as the indicator pathogen based on the assumption that A. suum is typically more difficult to kill than the eggs, cysts, and oocysts of most other parasites. The dairy cattle manure was piled into a biodrying storage shed. The forced aeration and natural decomposition processes heated a major portion of the waste pile to temperatures exceeding 55°C. The A. suum eggs were inoculated into special chambers and placed at three different elevations at intervals along the length of the pile. No viable eggs were recovered from any of the chambers removed from the compost pile. The complete inactivation of these helminths’ eggs by the process suggests that it would also be successful in the inactivation of oocysts. Compost of dairy cattle manure is a process used both on small farms and very large commercial applications. Composting, if performed well, can be very successful in the inactivation of all oocysts present in the treated piles. The temperatures reached in the process are fully capable of killing oocysts. The most important part of the composting process is monitoring to make certain it is turned appropriately and that adequate temperatures are reached throughout the pile.
D.
Oocysts in Manure Applied to Soil
An increased frequency in the spreading of dairy manure on pastures causes an increase in the risk of finding oocysts in the surface waters conjoining the areas where the manure is spread (Sischo et al., 2000). This indicates that the oocysts are not permanently bound into a matrix within the manure from which they cannot escape. Kuczysnska et al. (2005) found that oocysts in manure were more likely to attach to soil particles than oocysts that had been washed free of feces and were in water. However, when the concentration of the manure increased to greater than 0.1%, the effect was reversed and, again, the oocysts were less likely to attach to soil particle. Temperature did not affect the release rate of oocysts from soil (Schijven et al., 2004). However, oocysts from calf manure adhered more tightly to soil particles than did those from cow manure. Therefore, the mixing of cow and calf manure probably promotes a more efficient release of oocysts from the material. It was also found that the application of water in the form of drops rather than as mist increased oocyst release, as did an increased application rate. Thus, increased precipitation events on a farm are expected to increase the number of oocysts that leave the soil on which manure has been applied. Not only the type of manure in which oocysts are spread on pasture but the type of soil on the pasture affect the rate at which oocysts leave a pasture or move through the soil during a rain event. Oocysts adhered more tightly to soils with increased clay content (Kuczysnska et al., 2005), and oocysts in solution adhered to different types of particles, which affected their sedimentation rate (Searcy et al., 2005).
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VIII. Beef Cattle Manure and Treatment The environmental load of oocysts in feces from feedlot cattle in the central and western United States was examined by Atwill et al. (2006b). They found a point prevalence of around 1% with 1.3 to 3.6 oocysts per gram of feces based on an examination of 5274 fecal samples from 22 feedlots. This translates to 28,000 to 140,000 oocysts per steer per day; compared to dairy calves, this is a very low number. There are some 11 million steers in feedlots in the United States at any given time, producing around 330,000 t of feces each day; thus, there would be some 300 to 1500 billion oocysts leaving feedlots each day in the United States. The risk from feedlot cattle is markedly less than from dairy calves because the number of oocysts produced is markedly less per animal, but this study also showed that a portion of the recovered oocysts were C. parvum, so some concern over potential zoonotic disease from feedlots persists even though the risk appears very small.
A.
Composting Treatment
Feedlots, similar to large dairy operations or consortiums, often operate mammoth composting facilities to treat waste before disposal. It is assumed that in these large facilities with proper aeration and turning of manure piles, along with appropriate temperature monitoring, that the inactivation of oocysts and other pathogens is fairly routine. Inactivation of oocysts in beef feedlot manure by windrow composting during the summer and early fall in Lethbridge, Alberta, Canada, was examined by Van Herk et al. (2004). Manure piles to which barley straw was added took 23 days to reach 55°C, and piles with wood chips took 30 days to reach 55°C or greater. The piles were then maintained at or above these temperatures for an additional 30 days. Oocysts were rapidly inactivated once the temperature reached 55°C. All were nonviable in the piles with straw and wood chips after 42 and 56 days of composting, respectively.
B.
Runoff Control
A significant amount of work has been done on the effect of vegetated buffer strips on the removal of oocysts from surface waters (Atwill et al., 2006a). These buffers protect surface waters by causing the infiltration of overland flow into the soil, subsurface straining, and adsorption of the entrained pathogens. Runoff control is a useful barrier around all waste facilities that will aid in reducing the amount of environmental risk from a system if there is an overflow, flooding, or breakdown scenario. There are numerous means of providing excellent barriers of this type, and again, the reader is referred to the USDA’s Natural Resources Conservation Service Web site (http://www.nrcs.usda.gov).
IX. Swine Manure and Treatment Cryptosporidiosis in swine has only recently received attention as a separate group of disease-causing organisms. For several years, almost all infections in farm animals were considered to be due to a single species, C. parvum. With the application of molecular methods to identify oocysts from animals and humans, there are now 16 species and nearly 40 genotypes recognized within the genus Cryptosporidium (Fayer, 2007). In pigs, C. suis was found in 8 of 12 pigs from Australia (7 pigs) and Switzerland (1 pig) (Ryan et al., 2004) and has also been reported from pigs in Calgary, Canada. (Guselle et al., 2003). Pigs are also infected with a novel swine genotype, Cryptosporidium pig genotype II and, occasionally, C. parvum (Ryan et al., 2004). Cryptosporidium suis is not infectious to neonatal nude mice and poorly, if at all, to calves. It is less virulent but produces more oocysts in swine than does C. parvum. Cryptosporidium suis has been reported in only two humans (Xiao et al., 2002; Leoni et al., 2006). Farrow-to-finish swine agriculture and waste management involves four separate operations: (1) breeding and gestation, (2) farrowing sows, (3) nursery (getting piglets ready for finishing), and (4) finishing (feeding pigs from the nursery to market size of around 240 lb). Because of hygiene
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considerations, each of these stages requires a separate facility, including separate waste management operations. These waste management operations usually consist of a pit underneath the area where the swine are housed. The waste material in the underfloor pit is either drained by gravity or is pumped into a lagoon for anaerobic digestion using effluent from the waste lagoon as wash water to drain the pits. Solids and liquid are agitated to make a slurry before the waste in the pit is pumped to the storage lagoon. Often, water is added to the slurry to bring the solids content to less than 4%. The most common storage lagoon is an anaerobic lagoon, the dimensions of which are based on a certain cubic feet per pound of hog, with extra dimensions to account for flushing water, wash water, spillage, and precipitation. The primary lagoon may or may not drain into a secondary lagoon. Liquid from the lagoon is applied to agricultural fields by irrigation systems, usually a spray field.
A.
Lagoon Treatment
Waste lagoons, besides reducing the effluent BOD, also reduce the pathogen loads of the waste stream. Single-stage, primary swine waste lagoons substantially reduce concentrations of Salmonella, fecal coliforms, E. coli, enterococci, and coliphages in flushed swine waste, but do not reduce concentrations of C. perfrigens spores (Hill and Sobsey, 2003). The analysis of the effects of storing the oocysts of C. parvum from cattle in anaerobic pig slurries in England revealed about a 50% reduction in total oocyst numbers after 3 months, but oocysts still remained over 50% viable (Hutchison et al., 2005a) When these slurries were applied onto fescue plots, the number of oocysts declined over 1 to 2 months to undetectable levels, but there was no reduction in oocyst viability during this time, with all oocysts recovered being viable by the dye assay (Hutchison et al., 2005b).
B.
Anaerobic Digestion Treatment
Pigs are bred in very large numbers in Quebec and Ontario, Canada, where it was shown that the manure from these pigs contains oocysts of Cryptosporidium (Côté et al., 2006b). It is expected that mesophilic and thermophilic digestion of swine waste would have the same effects on oocysts as with other waste material. Because of the odors associated with swine manure and the cold climates in which many of the swine are grown in the world, psychrophilic anaerobic digesters (digesters run at lower temperatures, typically at 20°C or less) were examined for their ability to inactivate various pathogens, including oocysts (Côté et al., 2006a). Oocysts were present in 4 of the 20 manure slurries used to feed the reactors, and the oocysts were not detected after the digestion of the samples. It is unclear why the killed oocysts would be removed by this system with a retention time of 20 days, and it might be worthwhile to repeat the trial to examine inactivation and removal rates.
X.
Summary and Conclusions
Waste management is very important for the control of cryptosporidiosis. In the case of fecally transmitted pathogens, it would seem that the simplest means of prevention is to control environmental contamination with the infectious oocyst. Much of the importance of this organism as a disease agent is due to the oocyst’s ability to survive routine chlorination during sewage treatment of effluents and during treatment of drinking water. This resistance, the production of massive numbers of oocysts in naive and immunocompromised hosts, and resistance to treatment with a battery of anti-infectives are the major reasons that this pathogen has received such individual attention in recent years. However, as our knowledge has advanced, the threat has diminished. In the few years since the discovery of this organism as a major waterborne pathogen, great strides have been made in the disinfection of water. Barriers are present in sewage and drinking water systems that can remove or destroy oocysts before they can enter the environment or cause outbreaks from tap water. Advances in molecular genetics have made it possible to ascertain the true risks associated with the different species of Cryptosporidium as sources of human infections. At the same time, work has begun on preventing the spread of cryptosporidiosis on farms.
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Garcés G, Effengberger, M., Najdrowski, M., Wackwitz, C., Gronauer, A., Wilderer, P.A. and Lebuhn, M. 2006. Quantification of Cryptosporidium parvum in anaerobic digesters treating manure by (reversetranscription) quantitative realtime PCR, infectivity and excystation tests. Wat. Sci. Technol. 53, 195–202. Guselle, N.J., Appelbee, A.J. and Olson, M.E. 2003. Biology of Cryptosporidium parvum in pigs: from weaning to market. Vet. Parasitol. 113, 7–18. Hashimoto, A., Sugimoto, H., Morita, S. and Hirata, T. 2006. Genotyping of single Cryptosporidium oocysts in sewage by semi-nested PCR and direct sequencing. Wat. Res. 40, 2527–2532. Hill, V.R. and Sobsey, M.D. 2003. Performance of swine waste lagoons for removing Salmonella and enteric microbial indicators. Trans. Am. Soc. Agric. Eng. 46, 781–788. Hirata, T., Shimura, A., Morita, S., Suzuki, M., Motoyama, N., Hoshikawa, H., Moniwa, T. and Kaneko, M. 2001. The effect of temperature on the efficacy of ozonation for inactivating Cryptosporidium parvum oocysts. Wat. Sci. Technol. 43, 163–166. Hutchison, M.L., Walters, L.D., Moore, A. and Avery, S.M. 2005a. Declines of zoonotic agents in liquid livestock wastes stored in batches on-farm. J. Appl. Microbiol. 99, 58–65. Hutchison, M.L., Walters, L.D., Moore, T., Thomas, D.J.I. and Avery, S.M. 2005b. Fate of pathogens present in livestock wastes spread onto fescue plots. Appl. Environ. Microbiol. 71, 691–696. Jenkins, M.B., Anguish, L.J., Bowman, D.D., Walker, M.J. and Ghiorse, W.C. 1997. Assessment of dyepermeability assays for determination of inactivation rates of Cryptosporidium parvum oocysts. Appl. Environ. Microbiol. 63, 3844–3850. Jenkins, M.B., Walker, M.J., Bowman, D.D., Anthony, L.C. and Ghiorse, W.C. 1999. Use of a sentinel system for field measurements of Cryptosporidium parvum oocyst inactivation in soil and animal waste. Appl. Environ. Microbiol. 65, 1998–2005. Karpiscak, M.M., Freitas, R.J., Gerba, C.P., Sanchez, L.R. and Shamir, E. 1999. Management of dairy waste in the Sonoran desert using constructed wetland technology. Wat. Sci. Technol. 40, 57–65. Karpiscak, M.M., Sanchez, L.R., Freitas, R.J. and Gerba, C.P. 2001. Removal of bacterial indicators and pathogens from dairy wastewater by a multi-component treatment system. Wat. Sci. Technol. 44, 183–190. Kato, S., Fogarty, E. and Bowman, D.D. 2003. Effect of aerobic and anaerobic digestion on the viability of Cryptosporidium parvum oocysts and Ascaris suum eggs. Int. J. Environ. Hlth. Res. 13, 169–179. Kuczynska, E. Shelton, D.R. and Pachepsky, Y. 2005. Effect of bovine manure on Cryptosporidium parvum oocyst attachment to soil. Appl. Environ. Microbiol. 71, 6394–6397. Lemarchand, K. and Lebaron, P. 2003. Occurrence of Salmonella spp. and Cryptosporidium spp. in a French coastal watershed: relationship with fecal indicators. FEMS Microbiol. Lett. 218, 203–209. Leoni, F., Amar, C., Nichols, G., Pedraza-Diaz, S. and McLauchlin, J. 2006. Genetic analysis of Cryptosporidium from 2,414 humans with diarrhea in England between 1985 and 2000. J. Med. Microbiol. 55, 703–707. Li, H., Finch, G.R., Smith, D.W. and Belosevic, M. 2001. Chlorine dioxide inactivation of Cryptosporidium parvum in oxidant demand-free phosphate buffer. J. Environ. Eng. 127, 594–603. Logan, A.J., Stevik, T.R., Siegrist, R.L. and Rønn, R.M. 2001. Transport and fate of Cryptosporidium parvum oocysts in intermittent sand filters. Wat. Res. 35, 4359–4369. Lucio-Forster, A. 2006. Cryptosporidium genotypes recovered from wastewater effluents entering Lake Cayuga in the Cayuga Lake Watershed in upstate New York. In Detection and disinfection studies of microorganisms of health or religious concern in the agriculture and water industries: Ascaris suum, Cryptosporidium spp., and microcrustaceans. Ph.D. thesis, Cornell University, Ithaca, New York. Madore, M.S., Rose, J.B., Gerba, C.P., Arrowood, M.J. and Sterling, C.R. 1987. Occurrence of Cryptosporidium oocysts in sewage effluents and selected surface waters. J. Parasitol. 73, 702–705. Meunier, L., Canonica, S. and von Gunten, U. 2006. Implications of sequential use of UV and ozone for drinking water quality. Wat. Res. 40, 1864–1876. Montemayor, M., Valero, F., Jofre, J. and Lucena, F. 2005. Occurrence of Cryptosporidium spp. oocysts in raw and treated sewage and river water in north eastern Spain. J. Appl. Microbiol. 99, 1455–1462. Nieminski, E.C. and Ongerth, J.E. 1995. Removing Giardia and Cryptosporidium by conventional treatment and direct filtration. Am. Wat. Works Assoc. J. 87, 96–106. Payment, P., Plante, R. and Cejka, P. 2001. Removal of indicator bacteria, human enteric viruses, Giardia cysts, and Cryptosporidium oocysts at a large wastewater primary treatment facility. Can. J. Microbiol. 47, 188–193.
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Rimhanen-Finne, R., Vuorinen, A., MArmo, S., Malmberg, S. and Hänninen, M.-L. 2004. Comparative analysis of Cryptosporidium, Giardia and indicator bacteria during sewage sludge hygienization in various composting processes. Lett. Appl. Microbiol. 38, 301–305. Robertson, L.J., Hermansen, L. and Gjerde, B.K. 2006. Occurrence of Cryptosporidium oocysts and Giardia cysts in sewage in Norway. Appl. Environ. Microbiol. 72, 5297–5303. Robertson, L.J., Paton, C.A., Campbell, A.T., Smith, P.G., Jackson, M.H., Gilmour, R.A., Black, S.E., Stevenson, D.A. and Smith H.V. 2000. Giardia cysts and Cryptosporidium oocysts at sewage treatment works in Scotland, U.K. Wat. Res. 34, 2310–2322. Ryan, U.M., Monis, P., Enemark, H.L., Sulaiman, I., Samarasinghe, B., Read, C., Buddle. R., Robertson, I., Zhou, L., Thompson, R.C.A. and Xiao, L. 2004. Cryptosporidium suis n. sp (Apicomplexa: Cryptosporidiidae) in pigs (Sus scrofa). J. Parasitol. 90, 769–773. Schijven, J.F., Bradford, S.A. and Yang, S. 2004. Release of Cryptosporidium and Giardia from dairy cattle manure: physical factors. J. Environ. Qual. 33, 1499–1508. Searcy, K.E., Packman, A.I., Atwill, E.R. and Harter, T. 2005. Association of Cryptosporidium parvum with suspended particles: impact on oocyst sedimentation. Appl. Environ. Microbiol. 71, 1072–1078. Sischo, W.M., Atwill, E.R., Lanyon, L.E. and George, J. 2000. Cryptosporidia on dairy farms and the role these farms may have in contaminating surface water supplies in the northeastern United States. Prevent. Vet. Med. 43, 253–267. Smith, H.V., Grimason, A.M., Benton, C. and Parker, J.F.W. 1991. The occurrence of Cryptosporidium spp. oocysts in Scottish waters, and the development of a fluorogenic viability assay for individual Cryptosporidium spp. oocysts. Wat. Sci. Technol. 24, 169–172. Stadterman, K.L., Sninsky, A.M., Sykora, J.L. and Jakubowski, W. 1995. Removal and inactivation of Cryptosporidium oocysts by activated sludge treatment and anaerobic digestion. Wat. Sci. Technol. 31, 97–104. Stott, R., May, E., Matsushita, E. and Warren, A. 2001. Protozoan predation as a mechanism for the removal of Cryptosporidium oocysts from wastewaters in constructed wetlands. Wat. Sci. Technol. 44, 191–198. Stott, R., May, E., Ramirez, E. and Warren, A. 2003. Predation of Cryptosporidium oocysts by protozoa and rotifers: implications for water quality and public health. Wat. Sci. Technol. 47, 77–83. Suwa, M. and Suzuki, Y. 2003. Control of Cryptosporidium with wastewater treatment to prevent its proliferation in the water cycle. Wat. Sci. Technol. 47, 45–49. Timms, S., Slade, J.S. and Fricker, C.R. 1995. Removal of Cryptosporidium by slow sand filtration. Wat. Sci. Technol. 31, 81–84. Van Herk, F.H., McAllister, T.A., Cockwill, C.L., Guselle, N., Larney, F.J., Miller, J.J. and Olson, M.E. 2004. Inactivation of Giardia cysts and Cryptosporidium oocysts in beef feedlot manure by thermophilic windrow composting. Comp. Sci. Util. 12, 235–241. Villacorta-Martinez de Maturana, I., Ares-Mazás, M.E., Duran-Oreiro, D. and Lorenzo-Lorenzo, M.J. 1992. Efficacy of activated sludge in removing Cryptosporidium parvum oocysts from sewage. Appl. Environ. Microbiol. 58, 35145–1516. Ward, P.I., Deplazes, P., Regli, W., Rinder, H. and Mathis, A. 2002. Detection of eight Cryptosporidium genotypes in surface and waste waters in Europe. Parasitology 124, 359–368. Whitmore, T.N., and Robertson, L.J. 1995. The effect of sewage sludge treatment on oocysts of Cryptosporidium parvum. J. Appl. Bacteriol. 78, 34–38. Wohlsen, T., Bates, J., Gray, B., Aldridge, P., Stewart, S., Williams, M., and Katouli, M. 2006. The occurrence of Cryptosporidium and Giardia in the Lake Baroon catchment, Queensland, Australia. J. Wat. Supply, Res. Technol.—AQUA. 55, 357–366. Young, P.L. and Komisar, S.J. 2005. Settling behavior of unpurified Cryptosporidium oocysts in laboratory settling columns. Environ. Sci. Technol. 39, 2636–2644. Xiao, L., Bern, C., Arrowood, M., Sulaiman, I., Zhou, L., Kawai, V., Vivar, A., Lal, A.A., Gilma, R.H. 2002. Identification of the Cryptosporidium pig genotype in a human patient. J. Infect. Dis. 185, 1846–1848. Zuckerman, U., Gold, D., Shelef, G., and Armon, R. 1997. The presence of Giardia and Cryptosporidium in surface waters and effluents in Israel. Wat. Sci. Technol. 35, 381–384.
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14 Fish, Amphibians, and Reptiles
Thaddeus K. Graczyk
CONTENTS I.
Piscine Cryptosporidiosis............................................................................................................. 387 Cryptosporidium spp. in Fish ......................................................................................... 387 A. B. Transmission Studies....................................................................................................... 388 II. Amphibian Cryptosporidiosis ...................................................................................................... 388 Cryptosporidium spp. in Amphibians............................................................................. 388 A. B. Transmission Studies....................................................................................................... 389 III. Reptilian Cryptosporidiosis.......................................................................................................... 389 A. Clinical Signs and Pathology.......................................................................................... 389 B. Diagnosis of Cryptosporidium Infection in Reptiles ..................................................... 390 1. Barium.............................................................................................................. 390 2. Fecal Examination ........................................................................................... 390 3. Endoscopy ........................................................................................................ 390 4. Gastric Lavage and Cloacal Swabs ................................................................. 390 5. Regurgitated Material ...................................................................................... 390 6. Gastric Biopsies ............................................................................................... 391 7. Serum Antibody Test ....................................................................................... 391 8. Postmortem Examinations ............................................................................... 391 C. Treatment......................................................................................................................... 391 D. Prevention and Control ................................................................................................... 391 E. Transmission Studies....................................................................................................... 392 References........................................................................................................................... 392
I. A.
Piscine Cryptosporidiosis Cryptosporidium spp. in Fish
The species of Cryptosporidium infecting fish are C. molnari and C. scophthalmi (Alvarez-Pellitero and Sitja-Bobadilla, 2002; Alvarez-Pellitero et al., 2004). Merogonial and gamogonial stages of both species were in the typical extracytoplasmic position, whereas sporogonial stages were deep within the epithelium, with mature oocysts in parasitophorous vacuoles. The first report of Cryptosporidium infection in fish described a progressive illness lasting 2 months in a tropical marine fish, Naso lituratus (Hoover et al., 1981). Intestinal morphology was normal except for endogenous parasite stages in the microvillar surface. Although this parasite was named C. nasorum, no measurements or other taxonomically useful data were provided and, therefore, the name is considered a nomen nudum. Next, Cryptosporidium sp. was found as developmental stages in intestinal villi in 5 of 35 carp in Czechoslovakia (Pavlasek, 1983), and Cryptosporidium sp. oocysts were detected in intestinal contents of 5 brown trout from a reservoir near Sheffield, England (Rush et al., 1987). Neither the carp nor the trout were reported ill. Cryptosporidium sp. was also found in the stomach of hatchery-reared fry and fingerling cichlids from a lake in
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Israel (Paperna, 1983); red drum (Camus and Lopez, 1996) and barramundi (Lates calcacifier) in association with inflammatory cells in the intestinal lamina propria (Glazebrook and Campbell, 1987). Cryptosporidium molnari was described from two teleost fish, the gilthead sea bream (Sparus aurata) and the European sea bass (Dicentrarchus labrax) (Alvarez-Pellitero et al., 2002). The parasite was found mainly in the stomach epithelium and seldom in the intestine. External clinical signs consisted of whitish feces, abdominal swelling, and ascites. Histologically, the wide zones of epithelium invaded by oogonial and sporogonial stages appeared necrotic, with abundant cell debris and sloughing of epithelial cells into the lumen. Cryptosporidium molnari causes mortality in juvenile aquaculture-reared fish (Sitja-Bobadilla et al., 2005, 2006), and bacterial co-infection enhances the severity of the infection (Sitja-Bobadilla et al., 2006). Both C. molnari and C. scophthalmi have not been genetically characterized, which may present a problem in naming future piscine Cryptosporidium spp. However, histological, morphological, genetic, and phylogenetic analyses of a C. molnari-like isolate from a guppy (Poecilia reticulata) revealed that this parasite is genetically very distinct from all other species of Cryptosporidium (Ryan et al., 2004). Cryptosporidium scophthalmi was described from Scophthalmus maximus (Alvarez-Pellitero et al., 2004). The parasite was found mainly in the intestinal epithelium and very seldom in the stomach. External clinical signs were not detected, although some fish showed intestinal distension at necropsy (Alvarez-Pellitero et al., 2004). The marked histopathological damage in severe infections included distension of epithelial cells by large vacuoles containing clusters of oocysts, sloughing of epithelial cells, and detachment of intestinal mucosa (Alvarez-Pellitero et al., 2004). An inflammatory reaction involving leukocyte infiltration was sometimes observed.
B.
Transmission Studies
Although C. parvum oocysts from a human source were reported to be infectious for fish (Arcay et al., 1995), multiple attempts to experimentally infect 1.5-cm guppies (P. reticulata) and bluegills with C. parvum oocysts infectious for suckling mice were unsuccessful (Upton, 1990; Graczyk et al., 1996). Apparently, a slow passage of the inoculum oocyst in the fish gut or a prior undetected infection resulted in misdiagnosis of an experimental infection (Arcay et al., 1995). Attempts to infect rainbow trout (Oncorhynchus mykiss) with C. parvum oocysts also were unsuccessful (Freire-Santos et al., 1998). Cryptosporidium molnari was experimentally transmitted to gilthead sea bream (S. aurata) and European sea bass (D. labrax) by oral infection with infected stomach scrapings (Sitja-Bobadilla et al., 2003). The infection was also transmitted from infected gilthead sea bream to sea bass by cohabitation (SitjaBobadilla et al., 2003). Transmission of C. molnari is favored by cannibalism among cohabiting fish (Sitja-Bobadilla et al., 2003).
II. A.
Amphibian Cryptosporidiosis Cryptosporidium spp. in Amphibians
Reports on amphibian cryptosporidiosis are scant, and there are no valid species of Cryptosporidium described from amphibians. Feces collected from Ceratophrys ornata (Bell’s horned frog) at the Metropolitan Toronto Zoo were stained by modified Ziehl–Neelsen stain and revealed Cryptosporidium oocysts (Crawshaw and Mehren, 1987). Although oocysts were detected in the feces, the authors suggested they might have come from ingestion of infected mice (Crawshaw and Mehren, 1987). Cryptosporidium oocysts were incidentally detected in feces originating from American toad (Cranfield and Graczyk, 1995) and from hylidean frogs (Hassl, 1991). No information regarding pathology of these infections was provided in these reports. Cryptosporidiosis was reported in a 2-year-old emaciated female African clawed frog (Xenopus laevis) euthanized because of chronic weight loss (Green et al., 2003). There was no evidence of bacterial, fungal, or viral disease. A proliferative gastritis and the presence of numerous cryptosporidial stages throughout the intestine were observed in histologic specimens. Oocysts
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were present in water taken from the aquarium housing the infected frog and were likely shed by the sick frog. No attempt was made to identify the species of Cryptosporidium.
B.
Transmission Studies
One group of investigators reported experimental infection of amphibians with C. parvum oocysts of human origin (Arcay et al. 1995), whereas another group was unable to infect African clawed frogs (X. laevis) or poison dart frogs with C. parvum oocysts, which were infectious for suckling mice (Graczyk et al., 1996). Cryptosporidium serpentis oocysts obtained from six captive snakes were gastrically delivered to twelve Cryptosporidium-free African clawed frogs, nine tadpoles, and three recently metamorphosed adults of Cryptosporidium-free wood frogs (Graczyk et al., 1998). On days 7 and 14 post inoculation, no life cycle stages of Cryptosporidium were observed in any histological sections of stomach, jejunum, ileum, cloaca, or cecum. However, C. serpentis oocysts were recovered from the water in which the amphibians were kept, indicating that amphibians may disseminate C. serpentis oocysts in the natural habitat.
III. Reptilian Cryptosporidiosis Cryptosporidium serpentis described from snakes (Levine, 1980) and C. varanii (syn. C. saurophilum) described from lizards (Pavlasek et al., 1995; Koudela and Modry, 1998) are the only valid species, although as many as five species of Cryptosporidium infecting reptiles have been suggested based on oocyst morphology (Upton, 1990) and molecular studies (Xiao et al., 2004). Cryptosporidium serpentis infections in snakes and C. varanii infections in lizards are acquired for a lifetime, and self-cure was not observed (Cranfield and Graczyk, 1995, 2000, 2006; Cranfield et al., 1999). Although C. serpentis has been described from snakes, this species is virulent for lizards, and C. varanii described from lizards is virulent for snakes (Koudela and Modry, 1998; Xiao et al., 2004). Cryptosporidium serpentis preferentially infects the stomach, whereas C. varanii preferentially infects the intestine of snakes and lizards. Cryptosporidium has been reported in approximately 80 species of reptiles, including snakes, lizards, and tortoises (O’Donoghue, 1995; Fayer et al., 1997; Graczyk et al., 1998; Cranfield et al., 1999, 2000; Xiao et al., 2004; Cranfield and Graczyk, 2006). Most reports described clinical and subclinical infections in captive reptiles, whereas only subclinical infections have been reported in wild reptiles (Upton et al., 1989; Graczyk et al., 1997). Cryptosporidiosis is a common and sometimes life-threatening disease of captive snakes, lizards, and tortoises (Cranfield et al., 1999; Cranfield and Graczyk, 2000, 2006; Brownstein et al., 1977).
A.
Clinical Signs and Pathology
There are two manifestations of Cryptosporidium infections in reptiles: subclinical (i.e., carrier state), and clinical (i.e., gastritis, enteritis, and gastroenteritis) (Cranfield and Graczyk, 1995, 2000, 2006). Healthy reptiles are able to intermittently pass oocysts for years, oscillating between periods of excreting high numbers of oocysts to periods that are oocyst negative by acid-fast staining techniques. The prevalence of subclinically infected shedders can be high in a reptile collection (Carmel and Groves, 1993; Cranfield and Graczyk, 1995, 2000, 2006). Subclinical infections can be difficult to diagnosis because of the low oocyst output, sometimes far below the detection threshold of 3.75 × 104 oocysts/g (Cranfield and Graczyk, 2006; Graczyk et al., 1995, 1996), and intermittent patterns of oocyst voiding. Clinical signs in snakes are associated with gastric hyperplasia of the mucus-secreting cells (Brownstein et al., 1977). Snakes often have foul-smelling diarrhea and midbody swelling with reduction in lumen diameter; they can live from a few days up to 2 years after the appearance of clinical signs (Cranfield and Graczyk, 1995, 2000, 2006). Weight loss often occurs with persistent or periodical postprandial regurgitation 3 to 4 days after a meal. The gastric mucosa appears edematous with mucosal thickening and exaggerated longitudinal rugae that have copious amounts of mucus adhered to it. The
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surface may exhibit petechiae, enlarged rugae, excess mucous, and multiple foci of necrosis. Microscopy of gastric tissue reveals hyperplasia and hypertrophy of gastric glands, atrophy of granular cells, edema of the submucosa and lamina propria, and inflammation of the gastric mucosa characterized by infiltration with lymphocytes and heterophils. Cryptosporidium infections in lizards has been associated with acute enteritis and bacterial gastritis (Dillehay et al., 1986; Frost et al., 1994; Terrel et al., 2003; Taylor et al., 1999). Clinical signs include weight loss, anorexia, lethargy, and diarrhea. Histological examination usually reveals hyperplasia and mononuclear inflammation of the small intestine.
B. 1.
Diagnosis of Cryptosporidium Infection in Reptiles Barium
In snakes with postprandial regurgitation and midbody swelling, a barium study is useful to differentiate between gastric occlusion due to mucosal swelling and a nongastrointestinal mass (Cranfield and Graczyk, 1995, 2000, 2006).
2.
Fecal Examination
Historically, oocysts have been identified in reptile fecal specimens by examination of direct fecal smears stained with acid-fast stain. Epitopes of C. serpentis oocyst wall antigens produce positive reactions with fluorescein-labeled monoclonal antibodies that are commercially available. The MERIFLUORTM test is over 16 times more sensitive than acid-fast stain for detection of C. serpentis oocysts (Graczyk et al., 1995). However, even with the increased sensitivity of the immunofluorescent antibody (IFA), multiple negative fecal tests must be performed to raise the confidence level that a snake is truly negative for Cryptosporidium. Oocysts of Cryptosporidium mouse genotype are frequently seen in captive reptiles, which can confound the diagnosis of cryptosporidiosis (Xiao et al., 2004).
3.
Endoscopy
Endoscopy requires expensive equipment, and visual images of the gastric rugae and intestinal epithelium are difficult to interpret (Cranfield and Graczyk, 1995, 2000, 2006).
4.
Gastric Lavage and Cloacal Swabs
Gastric lavage and cloacal swabbing can be performed on inappetant and nondefecating reptiles (Graczyk et al., 1996). Cloacal swab smears were demonstrated to be far less effective than gastric lavage smears. Because the pathogen resides in the stomach area, it is expected that in nondefecating reptiles higher concentrations of oocysts would be found in stomach aspirates than in cloacal swabs. Gastric lavage is performed by passing a tube into the stomach located at the midpoint between the head and the cloaca of a snake. Phosphate-buffered saline (equal to 2% of total body weight) is passed through the tube into the stomach and then aspirated with the snake held head down, retrieving approximately 50% of the administered fluid. The aspirate is centrifuged and a smear prepared from the pellet. Stomach aspirates contain little particulate matter and, therefore, acid-fast stain detection is nearly as sensitive as IFA staining (Graczyk et al., 1996). Additionally, it was noted that the gastric lavage test was more sensitive if performed within 3 days of eating (Graczyk et al., 1996). Because the metabolic rate of gastric mucosal tissue increases over 22 times after a meal, it is proposed that the Cryptosporidium reproductive cycle increases with the metabolic increase in the gastric mucosa. It may be beneficial to administer an appropriate baby food via a stomach tube meal to an inappetant reptile 3 days prior to the stomach lavage.
5.
Regurgitated Material
Examination of smears of the parasite-rich mucus surrounding a regurgitated meal utilizing either the acid-fast or IFA stain can provide a definitive diagnosis (Cranfield and Graczyk, 1995, 2000, 2006).
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Gastric Biopsies
This is a relatively safe procedure that can aid in the prognosis of a case when Cryptosporidium developmental stages are found in the biopsy material. However, the nonuniform distribution of the pathogen on the gastric mucosa or intestinal epithelium makes a negative outcome difficult to interpret (Cranfield and Graczyk, 1995, 2000, 2006).
7.
Serum Antibody Test
Serum antibody enzyme-linked immunosorbent assay (ELISA) utilizing C. serpentis oocyst wall antigen has shown great sensitivity and specificity in surveys of reptile collections and in blind studies (Graczyk and Cranfield, 1997). The test diagnoses exposure to Cryptosporidium and can identify Cryptosporidiumnegative reptiles to establish pathogen-free colonies, and for selection for Cryptosporidium research.
8.
Postmortem Examinations
Several stomach and intestinal (for C. varanii) tissue sections should be obtained for histological examination (Cranfield and Graczyk, 1995, 2000, 2006). On rare occasions, Cryptosporidium-positive reptiles diagnosed on fecal samples were found to be negative on postmortem examination when limited samples of gastric tissue were collected.
C.
Treatment
Treatment regimes for reptiles have originated from the experiences of human and domestic animal treatment. The high morbidity and moderate mortality caused by Cryptosporidium in ophidian collections is due to the lack of anticryptosporidial pharmaceuticals that can be safely and efficaciously used for prophylaxis or therapy (Cranfield and Graczyk, 1995, 2000, 2006). For reptiles, trimethoprim sulfa (30 mg/kg) once a day for 14 days and then 1 to 3 times weekly for several months, spiramycin 160 mg/kg for 10 days, and paromomycin at 100 mg/kg for 7 days and then twice a week for 3 months has been used (Graczyk et al., 1996). Halofuginone, paromomycin, and spiramycin reduced the number of voided oocysts, but did not eliminate infection as determined by histological sections. Furthermore, halofuginone was severely hepatotoxic and nephrotoxic for snakes that were already physiologically debilitated by chronic Cryptosporidium infections. Trimethoprim-sulfamethoxazole and trimethoprim-sulfadiazine treatment resulted in oocyst-negative stools, but gastric biopsies of treated snakes revealed the pathogen in the mucosa (Cranfield and Graczyk, 1995; Graczyk et al., 1996). Therapy based on the protective passive immunity of hyperimmune bovine colostrum (raised against C. parvum in dairy cows immunized during gestation) was efficacious in treating subclinical and clinical C. serpentis infections in snakes and lizards (Graczyk et al., 1998, 1999, 2000). Six gastric colostrum treatments, delivered at 1% of the snake’s weight at weekly intervals histologically cleared C. serpentis in subclinical infections and regressed gastric histopathological changes. Supportive treatments, such as high temperatures, subcutaneous fluids, regular stomach tubing of highly digestible foods, and the elimination of any concurrent disease problems, appear to act synergistically with treatment aimed at Cryptosporidium.
D.
Prevention and Control
Cryptosporidium can be transmitted directly via the fecal-oral route or indirectly by contamination of food or water, e.g., utensils, feeding bottles, and cages (Cranfield and Graczyk, 1995, 2000, 2006). The oocysts, which are fully sporulated and infectious when excreted, are resistant to environmental stressors and to a wide range of commonly used disinfectants. Ammonia (5%) and formal saline (10%) were the most effective in altering oocysts’ infectivity after 18 h at > 4˚C (Cranfield and Graczyk, 1995). Strict high-standard hygiene, good management, and isolation of infected animals are essential in prevention of spreading of Cryptosporidium within captive reptiles.
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Cryptosporidium and Cryptosporidiosis, Second Edition Transmission Studies
Attempts by two groups to experimentally infect suckling mice with numerous isolates of oocysts of C. serpentis have failed (Tilley et al., 1990; Fayer et al., 1995). One group of investigators reported experimental infection of snakes with C. parvum oocysts of human origin (Arcay et al., 1995), whereas another group was unable to infect corn snakes with C. parvum oocysts of bovine origin that were infectious for suckling mice (Graczyk et al., 1996). Also, attempts to experimentally infect birds with C. serpentis have failed (Graczyk et al., 1998). Six 2-week-old Cryptosporidium-free Peking ducklings (Anas platyrhynchos) each received 2.0 × 106 viable C. serpentis oocysts from six naturally infected captive snakes. Histological sections of digestive (stomach, jejunum, ileum, cloaca, and cecum) and respiratory tract tissues (larynx, trachea, and lungs) did not contain life-cycle stages of Cryptosporidium in any of the inoculated ducklings (Graczyk et al., 1998). Under certain circumstances such as after the ingestion of C. serpentis-infected prey, herpetivorous birds might disseminate C. serpentis oocysts in the environment. In another experiment, groups of four to five, 3-month-old rat snakes (Elaphe obsoleta) were separately gastrically inoculated with 2.0 × 106 viable oocysts of C. muris (mice and calves), C. muris-like (Bactrian camels), C. wrairi (guinea pigs), C. baileyi (chickens), C. meleagridis (turkeys), Cryptosporidium sp. (turtles, tortoises, chameleons, and lizards), and C. serpentis from clinically (fatal case) and subclinically infected snakes (Graczyk and Cranfield, 1998). None of the snakes inoculated with oocysts originating from homothermous vertebrates developed infection as determined by histology and serology, whereas all snakes challenged with reptilian oocyst isolates were infected with Cryptosporidium on weeks 6 and 10 post inoculation. The study demonstrated that Cryptosporidium infections in snakes maintained on a diet of rodents or birds cannot be initiated via ingestion of an infected food item; however, snakes can void ingested oocysts, and housing snakes with other reptiles can enhance transmission of Cryptosporidium to snakes, and therefore should be avoided (Graczyk and Cranfield, 1998).
References Alvarez-Pellitero, P., Quiroga, M.I., Sitja-Bobadilla, A., Redondo, M.J., Palenzuela, O., Padros, F., Vazquez, S. and Nieto, J.M. 2004. Cryptosporidium scophthalmi n. sp. (Apicomplexa: Cryptosporidiidae) from cultured turbot Scophthalmus maximus. Light and electron microscope description and histopathological study. Dis. Aquat. Orgs. 62, 133–145. Alvarez-Pellitero, P. and Sitja-Bobadilla A. 2002. Cryptosporidium molnari n. sp. (Apicomplexa: Cryptosporidiidae) infecting two marine fish species, Sparus aurata L. and Dicentrarchus labrax L. Intl. J. Parasitol. 32, 1007–1021. Arcay, L., Baez de Borges, E. and Bruzual, E. 1995. Cryptosporidiosis experimental en la escala de vertebreados. I. Infecciones experimentales. II. Estudio histopatologico, Parasitol. al Dia, 19, 20–29. Brownstein, D., Strandberg, J.D., Montali, R.J., Bush, M. and Fortner, J. 1977. Cryptosporidium in snakes with hypertrophic gastritis. Vet. Pathol. 14, 606–617. Camus, A. and Lopez, A. 1996. Gastric cryptosporidiosis in juvenile Red Drum. J. Aquat. Anim. Hlth. 8, 167–172. Carmel, B. and Groves, V. 1993. Chronic cryptosporidiosis in Australian elapid snakes: control of an outbreak in a captive colony. Aust. Vet. J. 70, 293–295. Cranfield, M.R. and Graczyk, T.K. 1995. Cryptosporidiosis, in Manual of Reptile Medicine and Surgery, Mader D.R., Ed., W.B. Saunders Company, Philadelphia, PA, pp. 359–363. Cranfield, M.R. and Graczyk, T.K. 2000. Cryptosporidia in reptiles, in Kirk’s Current Veterinary Therapy XIII Small Animal Practice, Bonagura J.D., Ed., W.B. Saunders Company, Philadelphia, PA, pp. 1188–1191. Cranfield, M.R. and Graczyk, T.K. 2006. Cryptosporidiosis, in Manual of Reptile Medicine and Surgery, Mader, D.R., Ed., Saunders Elsevier, St. Louis, MO, pp. 756–762. Cranfield, M.R., Graczyk, T.K., Wright, K., Frye, F.L. and Raphael, B. 1999. Cryptosporidiosis. Bull. Assoc. Rept. Amph. Vet. 9, 15–21.
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Crawshaw, G.J. and Mehren, K.G. 1987. Cryptosporidiosis inzoo and wild animals, in Erkrankungen der Zootiere, Verhandlungsbericht des 29 Int. Symp. uber die Erkrankungen der Zootiere von 20, Ippen, R., and Schroder, H. D., Eds., Akad. Verlag, Berlin, pp. 353–362. Dillehay, D.I., Boosinger, T.R. and MacKenzie, S. 1986. Gastric cryptosporidiosis in a chameleon. J. Am. Vet. Med. Assoc. 189, 1139–1140. Fayer, R., Graczyk, T.K. and Cranfield, M.R. 1995. Multiple heterogenous isolates of Cryptosporidium serpentis from captive snakes are not transmissible to neonatal BALB/c mice (Mus musculus). J. Parasitol. 81, 482–484. Fayer, R., Speer, C.A. and Dubey, J.P. 1997. The general biology of Cryptosporidium, in Cryptosporidium and Cryptosporidiosis, Fayer, R., Ed., CRC Press, Boca Raton, FL, pp. 1–42. Freire-Santos, F., Vergara-Castiblanco, C.A., Tojo-Rodriguez, J.L., Santamarina-Fernandez, T. and Ares-Mazas, E. 1998. Cryptosporidium parvum: an attempt at experimental infection in rainbow trout Oncorhynchus mykiss. J. Parasitol. 84, 935–938. Frost, D., Nichols, D.K. and Citino, S.B. 1994. Gastric cryptosporidiosis in two ocellated lacertas (Lacerta lepida). J. Zoo Wildl. Med. 25, 138–142. Glazebrook, J.S. and Campbell, R.S.E. 1987. Diseases of barramundi (Lates calcarifer) in Australia: a review, in Management of Wild and Cultured Sea Bass/Barramundi (Lates calcarifer), Copland, J.W. and Gray, D.L., Eds., Australian Centre for International Agricultural Research, Canberra. pp. 204–206. Graczyk, T.K., Balazs, G.H., Work, T., Aguirre, A.A., Ellis, D.M., Murakawa, S.K.K. and Morris, R. 1997. Cryptosporidium sp. infections in green turtles, Chelonia mydas, as a potential source of marine waterborne oocysts in theHawaiian Islands. Appl. Environ. Microbiol. 63, 2925–2927. Graczyk, T.K. and Cranfield, M.R. 1996. Assessment of the conventional detection of fecal Cryptosporidium serpentis oocysts of subclinically infected captive snakes. Vet. Res. 27, 185–192. Graczyk, T.K. and Cranfield, M.R. 1997. Detection of Cryptosporidium-specific immunoglobulins in captive snakes by a polyclonal antibody in the indirect ELISA. Vet. Res. 28, 131–142. Graczyk, T.K. and Cranfield, M.R. 1998. Experimental transmission of Cryptosporidium oocyst isolates from mammals, birds, and reptiles to captive snakes. Vet. Res. 29, 187–195. Graczyk, T.K., Cranfield, M.R. and Bostwick, E.F. 1999. Hyperimmune bovine colostrum treatment of moribund Leopard geckos (Eublepharis macularius) infected with Cryptosporidium sp. Vet. Res. 30, 377–382. Graczyk, T.K., Cranfield, M.R. and Bostwick, E.F. 2000. Successful hyperimmune bovine colostrum treatment of Savanna monitors (Varanus exanthematicus) infected with Cryptosporidium sp. J. Parasitol. 86, 631–632. Graczyk, T.K., Cranfield, M.R. and Fayer, R. 1995. A comparative assessment of direct fluorescence antibody, modified acid fast stain, and sucrose flotation techniques for detection of Cryptosporidium serpentis oocysts in snake fecal specimens. J. Zoo Wildl. Med. 26, 396–402. Graczyk, T.K., Cranfield, M.R. and Fayer, R. 1998a. Oocysts of Cryptosporidium from snakes are not infectious to ducklings but retain viability after intestinal passage through a refractory host. Vet. Parasitol. 77, 33–40. Graczyk, T.K., Cranfield, M.R. and Geitner, A. 1998b. Multiple Cryptosporidium serpentis oocyst isolates from captive snakes are not transmissible to amphibians. J. Parasitol. 84, 1298–1300. Graczyk, T.K., Cranfield, M.R., Helmer, P., Fayer, R. and Bostwick, E.F. 1998c. Therapeutical efficacy of hyperimmune bovine colostrum treatment against clinical and subclinical Cryptosporidium serpentis infections in captive snakes. Vet. Parasitol. 74, 123–132. Graczyk, T.K., Cranfield, M.R., Mann, J. and Strandberg, J.D. 1998d. Intestinal Cryptosporidium sp. infection in Egyptian tortoise (Testudo kleinmanni). Intl. J. Parasitol. 28, 1885–1888. Graczyk, T.K., Cranfield, M.R. and Hill, S.L. 1996. Therapeutical efficacy of spiramycin and halofuginone treatment against Cryptosporidium serpentis (Apicomplexa: Cryptosporidiidae) infections in captive snakes. Parasitol. Res. 82, 143–148. Graczyk, T.K., Fayer, R. and Cranfield, M.R. 1996a.Cryptosporidium parvumis not transmissible to fish, amphibia, or reptiles. J. Parasitol. 82, 748–751. Graczyk, T.K., Owens, R. and Cranfield, M.R. 1996b. Diagnosis of subclinical cryptosporidiosis in captive snakes based on stomach lavage and cloacal sampling. Vet. Parasitol. 67, 143–151. Green, S.L., Bouley, D.M., Josling, C.A. and Fayer, R. 2003. Cryptosporidiosis associated with emaciation and proliferative gastritis in a laboratory-reared South African clawed frog (Xenopus laevis). Compar. Med. 53, 81–84.
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Hassl, A. 1991. An asymptomatic Cryptosporidia (Apicomplexa: Coccidia) infection in Agalychnis callidryas (COPE, 1862) (Anura: Hylidae). Herpetozoa 4, 127–131. Hoover, D.M., Hoerr, E.J., Carlton, W.W., Hinsman, E.J. and Ferguson, H.W. 1981. Enteric cryptosporidiosis in a Naso tang, Naso lituratus Bloch and Schneider. J. Fish Dis. 4, 425–428. Koudela, B. and Modry, D. 1998. New species of Cryptosporidium (Apicomplexa: Cryptosporidiidae) from lizards. Folia Parasitol. 45, 93–100. Levine, N.D. 1980. Some corrections of coccidian (Apicomplexa: Protozoa) nomenclature, J. Parasitol., 66, 830–834. O’Donoghue, P.J. 1995. Cryptosporidium and cryptosporidiosis in man and animals, Int. J. Parasitol. 25, 139–195. Paperna, I. 1987. Scanning electron microscopy of the coccidian parasite Cryptosporidium sp. from cichlid fishes. Dis. Aquat. Org. 3, 231–232. Pavlasek, I. 1983. Cryptosporidium sp. in Cyprinus carpio Linneus 1758 in Czechoslovakia. Folia Parasitol. 30, 248–251. Pavlasek, I., Lavickova, M., Horak, P., Kral, J. and Kral, B. 1995. Cryptosporidium varanii n. sp. (Apicomplexa, Cryptosporidiidae) in Emerald monitor (Varanus prasinus) Schegel 1893) in captivity at Prague zoo. Gazella 22, 99–108. Rush, B.A., Chapman, P.A. and Ineson, R.W. 1987. Cryptosporidium and drinking water. Lancet, 2, 632–633. Ryan, U., O’Hara, A. and Xiao, L. 2004. Molecular and biological characterization of a Cryptosporidium molnari-like isolate from a guppy (Poecilia reticulata). Appl. Environ. Microbiol. 70, 3761–3765. Sitja-Bobadilla, A. and Alvarez-Pellitero, P. 2003. Experimental transmission of Cryptosporidium molnari (Apicomplexa: Coccidia) to gilthead sea bream (Sparus aurata L.) and European sea bass (Dicentrarchus labrax L.). Parasitol. Res. 91, 209–214. Sitja-Bobadilla, A., Padros, F., Aguilera, C. and Alvarez-Pellitero, P. 2005. Epidemiology of Cryptosporidium molnari in Spanish gilthead sea bream (Sparus aurata L.) and European sea bass (Dicentrarchus labrax L.) cultures: from hatchery to market size. Appl. Environ. Microbiol. 71, 131–139. Sitja-Bobadilla, A., Pujalte, M.J., Macian, M.C., Pascual, J. and Alvarez-Pellitero, P. 2006. Interactions between bacteria and Cryptosporidium molnari in gilthead sea bream (Sparus aurata) under farm and laboratory conditions. Vet. Parasitol. Epub ahead of print. Taylor, M.A., Geach, M.R. and Cooley, W.A. 1999. Clinical and pathological observations on natural infections of cryptosporidiosis and flagellate protozoa in leopard geckos (Eublepharis macularius). Vet. Rec. 145, 695–699. Terrell, S.P., Uhl, E.W. and Funk, R.S. 2003. Proliferative enteritis in leopard geckos (Eublepharis macularius) associated with Cryptosporidium sp. infection. J. Zoo Wildl. Med. 34, 69–75. Tilley, M., Upton, S.J. and Freed, P.S. 1990. A comparative study of the biology of Cryptosporidium serpentis and Cryptosporidium parvum (Apicomplexa: Cryptosporidiidae) J. Zoo Wildl. Med. 21, 463–467. Upton, S.J. 1990. Cryptosporidium spp. in lower vertebrates, in Cryptosporidiosis of Man and Animals, Dubey, J.P., Speer, C.A., and Fayer, R., Eds., CRC Press, Boca Raton, FL, pp.149–156. Upton, S.J., McAllister, C.T, Freed, P.S. and Barnard, S.M. 1989. Cryptosporidium spp. in wild and captive reptiles. J. Wildl. Dis. 25, 20–30. Xiao, L., Ryan, R.M., Graczyk, T.K., Limor, J., Li, L., Koumbert, M., Hunge, R., Sulaiman, I.M., Zhou, L., Arrowood, M J., Koudela, B., Modry, D. and Lal, A.A. 2004. Genetic diversity of Cryptosporidium spp. in captive reptiles. Appl. Environ. Microbiol. 70, 891–899.
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15 Birds*
Una M. Ryan and Lihua Xiao
CONTENTS I. II. III. IV. V. VI.
Introduction .................................................................................................................................. 395 Avian Cryptosporidiosis ............................................................................................................... 396 Immunity ...................................................................................................................................... 397 In Vitro and In Ovo Culture ......................................................................................................... 398 Chemoprophylaxis and Control ................................................................................................... 399 Cryptosporidium meleagridis Slavin, 1955 ................................................................................. 399 A. Biology ............................................................................................................................ 399 B. Life Cycle........................................................................................................................ 400 C. Pathogenesis .................................................................................................................... 400 D. Host Range ...................................................................................................................... 401 E. Genetic Analysis.............................................................................................................. 403 VII. Cryptosporidium baileyi Current, Upton and Haynes, 1986....................................................... 403 A. Biology ............................................................................................................................ 403 B. Life Cycle........................................................................................................................ 403 C. Pathogenesis .................................................................................................................... 406 D. Host Range ...................................................................................................................... 406 E. Genetic Analysis.............................................................................................................. 407 VIII. Cryptosporidium galli Pavlásek, 1999......................................................................................... 407 A. Biology ............................................................................................................................ 407 B. Life Cycle........................................................................................................................ 409 C. Pathogenesis .................................................................................................................... 409 D. Host Range ...................................................................................................................... 410 E. Genetic Analysis.............................................................................................................. 410 IX. Avian Genotypes .......................................................................................................................... 410 A. Avian Genotypes I–IV .................................................................................................... 410 B. Eurasian Woodcock Genotype ........................................................................................ 412 C. Duck Genotype................................................................................................................ 412 D. Goose Genotypes I–IV.................................................................................................... 412 References........................................................................................................................... 413
I.
Introduction
Cryptosporidiosis is one of the most prevalent parasitic infections in domesticated, caged, and wild birds (O’Donoghue, 1995; Sreter and Varga, 2000), and the parasite has been reported in more than 30 avian species worldwide, belonging to orders Anseriformes, Charadriiformes, Columbiformes, Galliformes, *
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.
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Passeriformes, Psittaciformes, and Struthioniformes (Lindsay and Blagburn, 1990; O’Donoghue, 1995; Fayer et al., 1997; Sreter and Varga, 2000; Ng et al., 2006). However, few studies have examined the genetic diversity of Cryptosporidium spp. among avian hosts. Only three avian Cryptosporidium spp. are recognized: Cryptosporidium meleagridis, Cryptosporidium baileyi, and Cryptosporidium galli. These three Cryptosporidium spp. can each infect numerous birds, but they differ in host range and predilection sites. Even though both C. meleagridis and C. baileyi are found in the small and large intestine and bursa, they differ significantly in oocyst size, and only C. baileyi is also found in the respiratory tissues such as the conjunctiva, sinus, and trachea. In contrast, C. galli infects only the roventriculus. Two other species of Cryptosporidium have been named from birds: Cryptosporidium anserinum from a domestic goose (Anser domesticus) (Proctor and Kemp, 1974) and Cryptosporidium tyzzeri from chickens (Gallus gallus domesticus) (Levine, 1961). Neither of these reports gave adequate descriptions of oocysts or provided other useful information, and hence they are not considered valid species (Lindsay and Blagburn, 1990; Chapter 1). A number of genetically distinct avian genotypes have, however, been recently described, which are likely to be categorized as species in the future once more biological information becomes available.
II.
Avian Cryptosporidiosis
Naturally occurring cryptosporidiosis in birds manifests itself in three clinical forms: respiratory disease, enteritis, and renal disease. Usually only one form of the disease is present in an outbreak (Lindsay and Blagburn, 1990). Although three Cryptosporidium species are considered to be valid in birds and a number of avian genotypes have been identified, most reports of natural infections have not provided enough information to conclusively determine the species or genotype involved. It is therefore difficult to say with certainty which species or genotype is responsible for which form of avian cryptosporidiosis. Cryptosporidium meleagridis was originally described from the intestines of turkeys (Meleagris gallopavo) (Slavin, 1955) and is thought to be more frequently associated with enteritis (Lindsay and Blagburn, 1990). Cryptosporidium baileyi was first described from the bursa, cloaca, and respiratory tract of chickens (Current et al., 1986) and is more frequently associated with respiratory cryptosporidiosis, whereas C. galli has been described from the proventriculus only (Pavlásek, 1999; Ryan et al., 2003b). Overlap has been reported between C. meleagridis and C. baileyi in their site of infection (Lindsay and Blagburn, 1990; O’Donoghue, 1995), but to date not enough is known about C. galli to determine if it can infect sites other than the proventriculus of birds. Respiratory disease is the most common form of cryptosporidiosis in chickens (Dhillon et al., 1981; Itakua et al., 1984; Goodwin et al., 1988a; Latimer et al., 1988; Nakamura and Abe, 1988; Fernandez et al., 1990). Only occasionally has renal (Nakamura and Abe, 1988) or small-intestinal involvement been reported in chickens (Gharagozlou and Khodashenas, 1985). Respiratory disease has also been reported in turkeys (Hoerr et al., 1978; Glisson et al., 1984; Gharagozlou and Khodashenas, 1985; Tarwid et al., 1985; Ranck and Hoerr, 1987), common quail (Coturnix coturnix) (Tham et al., 1982; O’Donoghue et al., 1987; Murakami et al., 2002), ring-necked pheasants (Phasianus colchicus) (Whittington and Wilson, 1985; O’Donoghue et al., 1987), and budgerigars (Melopsittacus undulatus) (O’Donoghue et al., 1987). Clinical signs are generally similar in all avian species and consist of rales, coughing, sneezing, and dyspnea (Lindsay and Blagburn, 1990). Excess mucus in the trachea and nasal cavities and airsacculitis may be present. Microscopic lesions generally consist of hypertrophy and hyperplasia of infected epithelial cells (Lindsay and Blagburn, 1990). Even though species identification was not undertaken in most of the reports, C. baileyi was likely the cause of the clinical signs and illness. Enteritis associated with Cryptosporidium has been reported in turkeys (Slavin, 1955; Goodwin et al., 1988b; Wages and Ficken, 1989; Gharagozlou et al., 2006), bobwhite quail (Colinus virginianus) (Hoerr et al., 1986; Guy et al., 1987), budgerigars (O’Donoghue et al., 1987; Ley et al., 1988; Goodwin and Krabill, 1989), cockatiels (Nymphicus hollandicus) (Goodwin and Krabill, 1989; Lindsay and Blagburn, 1990; Lindsay et al., 1991), a ring-necked parrot (Psittacula krameri) (Morgan et al., 2000b), a macaw, tundra swan (Cygnus columbianus) (Ley et al., 1988), and lovebirds (Agapornis spp.) (Belton and Powell,
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1987). High morbidity and mortality were associated with disease in bobwhite quail and lovebirds. Mildto-severe disease has been reported in turkeys (Slavin, 1955; Goodwin et al., 1988b; Wages and Ficken, 1989; Gharagozlou et al., 2006). Microscopic lesions consist of villous atrophy, villous fusion, and crypt hyperplasia. The villi were moderately atrophic, the crypts were hypertrophic, and the lamina propria was infiltrated by large numbers of lymphocytes, heterophils, and fewer macrophages and plasma cells. Numerous intraepithelial leukocytes and exocytosing inflammatory cells also were present (Goodwin et al., 1988b). In a recent study in Iran, naturally occurring cryptosporidiosis was reported in turkey poults suffering from diarrhea and unthriftiness. Histological and ultrastructural studies revealed a high number of Cryptosporidium developmental stages mainly located in the mid and terminal portions of the small intestine of the poults. Other portions of the intestinal tract were less frequently infected. Oocyst shedding was detected only in 29% of the histologically positive birds. Although genotyping was not performed, based on host species, clinical signs, pathology, and tissue location of the parasites, it was assumed that C. meleagridis was responsible for the infection (Gharagozlou et al., 2006). Renal cryptosporidiosis has been reported in black-throated finches (Poephila cincta) (Gardiner and Imes, 1984), jungle fowl (Gallus spp.) (Randall, 1986), and chickens (Nakamura and Abe, 1988). Kidneys were reported to be pale and enlarged and, occasionally, urate crystals were seen in surface tubules (Lindsay and Blagburn, 1990). The epithelial cells of the collecting ducts, collecting tubules, distal convoluted tubules, and ureters become hypertrophic and hyperplastic (Lindsay and Blagburn, 1990). Renal cryptosporidiosis was experimentally induced in specific-pathogen-free (SPF) chickens coinfected with C. baileyi and Marek’s disease virus (MDV). The kidneys were markedly swollen and pale, with visible urate crystals in the ureters and surface tubules, and oocysts of C. baileyi were confirmed in three cases by histological examination of paraffin-embedded kidney sections (Abbassi et al., 1999). Recently urinary tract cryptosporidiosis was reported in adult egg-laying chickens in the United States (Trampel et al., 2000). Numerous developing stages of Cryptosporidium were observed on the apical surface of epithelial cells lining renal collecting tubules and ureters. The species of Cryptosporidium was not confirmed. This is the first report of urinary tract cryptosporidiosis occurring in adult hens in a modern commercial egg production facility (Trampel et al., 2000).
III. Immunity Age-related resistance to clinical disease and the ability to clear orally or intratracheally induced C. baileyi infections have been examined in chickens ranging in age from 2 days to 9 weeks (Lindsay et al., 1988a; Taylor et al., 1994; Sreter et al., 1995). Results revealed that the prepatent period was significantly shorter and the patency significantly longer in younger birds. Clinical respiratory disease occurred only in birds up to 2 weeks old. Chickens infected at 1 week of age excreted thrice the number of oocysts excreted by those inoculated at 9 weeks of age. These results indicate that innate resistance to C. baileyi is age related. It is thought that the older birds have better-functioning immune systems and are therefore better able to mount an effective immune response and clear the parasites (Lindsay and Blagburn, 1990). Chickens that were orally inoculated with C. baileyi oocysts and cleared the infection were resistant to secondary oral inoculations (Current and Snyder, 1988). These birds passed no oocysts and had no parasites in their tissues. Serum antibodies to oocyst/sporozoite antigens were found as early as 7 days post inoculation (PI) (Current and Snyder, 1988). The significance of the serum antibody response is unclear. However, a more recent study suggested that serum antibodies play little role in acquired resistance to challenge infection (Hornok et al., 1996). Results of that study revealed that there was no significant difference between the antibody responses of birds challenged orally with 8 × 105 C. baileyi oocysts at the age of 4 weeks that had previously been either (1) immunized with C. baileyi oocystderived proteins injected intramuscularly or (2) inoculated orally with 8 × 105 viable C. baileyi at 2 weeks of age (Hornok et al., 1996). Attempts to produce in ovo vaccination with C. baileyi oocyst extracts were also unsuccessful (Hornok et al., 2000). Another study examined the correlation of circulating antibody and cell-mediated immunity (CMI) to resistance to C. baileyi using hormonal and chemical bursectomy in the one experiment and cyclosporin A (a calcineurin inhibitor that blocks T-cell
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activation) in a second experiment. In experiment 1, there was no correlation between antibody (determined by enzyme-linked immunosorbent assay) and resistance to infection as measured by body weight, gross lesions, morbidity, and mortality. Bursectomy altered antibody production, but not CMI, as measured by the delayed-type hypersensitivity skin reaction. In experiment 2, cyclosporin A reduced CMI, but not antibody production. Chicks treated with cyclosporin A were more susceptible to C. baileyi (more severe respiratory disease) than untreated controls. Results suggested that CMI is more important in resistance to C. baileyi than circulating antibody (Hatkin et al., 1993). Another study that examined the effects exerted by human recombinant interleukin-1 beta and the prostaglandin inhibitor indomethacin on the course of C. baileyi infection in chickens concluded that the systemic application of interleukin1 induced only partial protection against C. baileyi in chickens and that prolonged, abundant oocyst shedding is due to an indomethacin-sensitive immunodepression via the prostaglandin pathway (Hornok et al., 1999). The effect of an existing 7-day-old C. baileyi infection on the ability of chickens to produce an antibody response to vaccination with Newcastle disease virus (NDV) and infectious bursal disease virus (IBDV) and their ability to mount a delayed hypersensitivity reaction (DH) to avian oil tuberculin has been examined (Blagburn et al., 1987). No significant differences were seen in IBDV antibody titers compared to controls. However, significantly increased NDV antibody titers and decreased DH reactions were seen in chickens that were infected with C. baileyi. Experimental coinfections of C. baileyi and chicken anemia virus (CAV) suggest that concurrent CAV infection increases the reproductive potential of C. baileyi in chickens and both pathogens have a synergistic effect on each other (Hornok et al., 1998). Similarly, experimental coinfection of chickens with C. baileyi and Marek’s disease virus (MDV) showed a considerable synergistic effect in concurrently infected chickens and more severe consequences when chickens received MDV before C. baileyi infection (Abbassi et al., 2000). Coinfection with both pathogens induced more lasting oocyst shedding and severe clinical cryptosporidiosis with weakness, anorexia, depression, growth retardation, and chronic and severe respiratory disease leading to death in the majority of chickens (Abbassi et al., 2000). Coinfection of C. baileyi and MDV has also been shown to result in renal cryptosporidiosis (Abbassi et al., 1999). Little is known about resistance or immunity to C. meleagridis or C. galli infections.
IV.
In Vitro and In Ovo Culture
Little research has been conducted on the cell culture of avian Cryptosporidium species. Excystation of C. baileyi and C. meleagridis has been described in excystation solutions containing 0.75% sodium taurocholate (ST) in Hanks’ balanced salt solution (HBSS) or 0.75% ST plus 0.25% trypsin in HBSS (Sundermann et al., 1987a). Sporozoites of C. baileyi did not undergo development in primary cell cultures from either avian or mammalian hosts, or in mammalian cell lines (Lindsay et al., 1988b). Recently, successful cultivation of C. meleagridis from a human patient has been accomplished on Madin-Darby bovine kidney cells (Akiyoshi et al., 2003). Monolayers grown in 96-well microtiter plates were infected with various oocyst doses per well ranging from 2.5 × 104 to 2 × 105. After 48 h of incubation at 37°C, the cells were fixed and reacted with specific anti-C. parvum rabbit serum and the number of immunofluorescent-labeled parasites quantitated and compared with a C. parvum infection (Akiyoshi et al., 2003). Results revealed that C. meleagridis is capable of infecting mammalian cells and although the infection rate was lower for C. meleagridis than C. parvum at the lower doses, at the highest concentration, the rate of infection between the two species was comparable (Akiyoshi et al., 2003). No studies have been conducted to determine if C. galli will develop in culture. Complete development of C. baileyi has been described from the chorioallantoic membrane (CAM) of chicken embryos when sporozoites or oocysts were inoculated into the allantoic cavity (Lindsay et al., 1988b). Oocysts obtained after C. baileyi had been passaged up to 20 times in chicken embryos still caused clinical respiratory disease and gross air sacculitis when inoculated IT into 2-day-old broiler chickens. Oocysts that had been passaged 10 times in chicken embryos were similarly pathogenic for 4-day-old turkeys after intratracheal (IT) inoculation (Lindsay et al., 1988b). Complete development of C. baileyi in the CAM of turkey, domestic duck, muscovy duck (Cairina moschata), chukar partridge
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(Alectoris chukar), guinea fowl, ring-necked pheasant, bobwhite quail, and common (Japanese) quail (Coturnix japonica) embryos was also achieved (Lindsay et al., 1988b). One study compared the rate of reproduction of C. baileyi in eggs to that in chickens and concluded that although the rate of reproduction of C. baileyi seen in the eggs was only about 50% of that observed in chickens (~77 million per egg compared to 161 to 168 million from chickens), nearly the same number of oocysts could be obtained from two eggs as compared with one chicken (Wunderlin et al., 1997). No attempts have been made to examine the in ovo development of C. meleagridis and C. galli.
V.
Chemoprophylaxis and Control
At present no effective chemotherapy is available for the treatment of avian cryptosporidiosis. Experimental studies have shown that most commonly used anticoccidials when used alone or in combination with the dihydroquinoline antioxidant duokvin do not prevent or reduce respiratory disease in chickens inoculated with C. baileyi oocysts (Lindsay and Blagburn, 1990; Varga et al., 1995). Oxytetracycline, amprolium and chlortetracycline had no effect on respiratory cryptosporidiosis in turkeys and peafowl chicks (Mason and Hartley, 1980; Glisson et al., 1984). Similarly, oxytetracycline, neomycin, and furazolidone had no effect on intestinal cryptosporidiosis in bobwhite quail (Hoerr et al., 1986). Halofuginone, salinomycin, lasalocid, and monensin have also been evaluated but did not protect chickens from C. baileyi infection (Lindsay et al., 1987b). Another study examined the effects exerted by human recombinant interleukin-1 beta (hrIL-1 beta) and the prostaglandin inhibitor indomethacin on the course of C. baileyi infection in chickens (Hornok et al., 1999). Parenteral application of hrIL-1 beta decreased oocyst shedding to 6%, but the infection ran a similar course in treated and control birds. However, indomethacin mixed with feed lessened oocyst shedding to 13.7% and also shortened its duration (Hornok et al., 1999). A more recent study on the anticryptosporidial prophylactic efficacy of two commercially available antibiotics, enrofloxacin and paromomycin, reported that the efficacy of enrofloxacin was 52% at the recommended level and the efficacy of paromomycin was 67 to 82% (Sreter et al., 2002). This is the highest reported efficacy of all drugs tested against avian cryptosporidiosis to date (Sreter et al., 2002). Control methods against avian cryptosporidiosis generally rely on limiting or preventing infection. Most commonly used disinfectants are not effective when used at concentrations recommended by manufacturers (Sundermann et al., 1987b). Commercially available ammonia compounds when used at 50% (v/v) were effective, with < 5% of oocysts remaining viable. A similar concentration of commercial bleach (5.25% sodium hypochlorite) was somewhat effective, with <15% of oocysts remaining viable (Sundermann et al., 1987b).
VI. Cryptosporidium meleagridis Slavin, 1955 A.
Biology
Tyzzer first described avian cryptosporidiosis in 1929, when he reported that a species structurally similar to Cryptosporidium parvum found in mice was present in the ceca of chickens (Tyzzer, 1929). He did not name the organism or describe the oocysts, and it was not until 35 years later, in 1955, that Slavin found a structurally similar parasite in the ileum of turkey poults (Slavin, 1955). Based on dried smears stained by the MacNeal-modified Romanowsky method, the cytology and measurements of merozoites, trophozoites, meronts, gametes, and oocysts were determined, and the parasite was named C. meleagridis (Figure 15.1). Oval oocysts measuring 4.5 × 4.0 µm appeared indistinguishable from those of C. parvum (Slavin, 1955). Lindsay et al. (1989a) gave measurements of 5.2 × 4.6 (4.5 – 6.0 × 4.2 5.3) µm for viable oocysts from turkey feces (Table 15.1). Although enormous numbers were observed in smears, sporozoites could not be identified within them (Slavin, 1955). Viable oocysts measured 5.1 × 4.5 µm (Slavin, 1955).
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FIGURE 15.1 Nomarski Interference microscopy of Cryptosporidium meleagridis oocysts. Bar, 5 µm.
TABLE 15.1 Morphometric Comparisons of the Oocysts from Cryptosporidium meleagridis, Cryptosporidium baileyi, Cryptosporidium galli, and Avian Cryptosporidium Genotypes Oocyst Length (µm)
Oocyst Width (µm)
C. meleagridis C. baileyi C. galli Avian genotype IIa
4.5–6.0 6.0–7.5 8.0–8.5 6.0–6.5
4.2–5.3 4.8–5.7 6.2–6.4 4.8–6.6
Avian genotype III Avian genotype IV Eurasian woodcock genotype
7.5 8.25 8.5
6.0 6.3 6.4
Species/Genotype
a
B.
Length/Width Ratio 1.00–1.33 1.05–1.79 1.3 1–1.25 1.25 1.30 1.32
Reference Lindsay et al., 1989a Lindsay et al., 1989a Ryan et al., 2003b Meireles et al., 2006; Ng et al., 2006 Ng et al., 2006 Ng et al., 2006 Ryan et al., 2003a
Morphometric measurements are not available for additional avian genotypes listed in Chapter 1, Table 1.3.
Life Cycle
Little is known of the life cycle of C. meleagridis. Both asexual and sexual stages were described by Slavin in 1955 (Table 15.2). More recently, the life cycle of what is presumed to be C. meleagridis has been described from naturally infected 30-day-old commercial turkeys (Tacconi et al., 2001). Tissue section examinations revealed the presence the parasite in multiple organs, but its prevalence was highest in the intestinal and bursal epithelial cells (Tacconi et al., 2001). Endogenous developmental stages of a C. meleagridis isolate of human origin have also been described in a mouse and a chicken experimentally infected with this isolate (Figure 15.2) (Akiyoshi et al., 2003).
C.
Pathogenesis
The factors that influence the establishment or site of development of C. meleagridis infections have not been studied in detail. Oral inoculation of poults with C. meleagridis oocysts produced infections in the ileum, ceca, colon, cloaca, and bursa of Fabricius (BF) (Bermudez et al., 1988). Mucosal changes in chicken poults infected with C. meleagridis included a slight shortening of villi and an irregular
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401 TABLE 15.2 Comparison of the Developmental Stages of Cryptosporidium meleagridis and Cryptosporidium baileyi Developmental Stage
C. meleagridisa (µm)
First-generation meront First-generation merozoite Number of merozoites Second-generation meront Second-generation merozoites Number of merozoites Third-generation meront Third-generation merozoites Number of merozoites Microgamont Microgametes Number of microgametes Macrogamont Unsporulated oocyst Sporulated oocyst
5.0 × 4.0 5.0 × 1.1 8 Not seen Not seen Not seen Not seen Not seen Not seen 4.0 × 4.0 1.0 × 0.3 16 5.0 × 4.0 4.5 × 4.0 Not seen
a
b
C. baileyib (µm) 5.0 6.9 8 5.1 5.0 4 5.2 3.6 8 4.0 1.7 16 5.8 6.3 6.2
× 4.9 × 1.1 × 5.1 × 1.3 × 5.1 × 1.1 × 4.0 × 0.5 × 5.8 × 5.2 × 4.6
From Slavin, D. 1955. Cryptosporidium meleagridis (sp. nov.). J. Comp. Pathol. 65, 262–266. With permission. From Current, W.L., Upton, S.J. and Haynes, T.B. 1986. The life cycle of Cryptosporidium baileyi n. sp. (Apicomplexa, Cryptosporidiidae) infecting chickens. J. Protozool. 33, 289–296. With permission.
epithelial surface layer (Akiyoshi et al., 2003). Cryptosporidium meleagridis infection has been associated with diarrhea and unthriftiness (Slavin, 1955; Gharagozlou et al., 2006). Broiler chickens orally infected with 5 × 105 oocysts of C. meleagridis at 7, 14, and 21 days of age had apparent clinical signs of infection, but neither final live weight nor mortality was significantly influenced (Tumova et al., 2002). Growth retardation was observed in the 2-week period after infection. However, compensatory growth started when oocyst shedding ceased. The number of oocysts in fecal specimens of chickens infected with C. meleagridis was two to three times lower than in fecal specimens of chickens infected with C. baileyi. Chickens infected with both C. baileyi and C. meleagridis (with 5 × 105 oocysts of each) had significantly lower final live weight and reduced feed efficiency than chickens infected with C. baileyi alone. Concurrent infection did not influence individual C. baileyi or C. meleagridis oocyst shedding (Tumova et al., 2002).
D.
Host Range
Similar to C. parvum, C. meleagridis appears to have a large host range and has been found in a wide range of avian species including turkeys, parrots, chickens, and cockatiels (O’Donoghue, 1995; Morgan et al., 2000b; Sreter and Varga, 2000; Darabus and Olariu, 2003; Abe and Iseki, 2004). Experimental transmission studies have shown that C. meleagridis can infect broiler chickens, ducks, turkeys, calves, pigs, rabbits, rats, and mice (O’Donoghue, 1995; Akiyoshi et al., 2003; Darabus and Olariu, 2003; Huang et al., 2003). The distribution of the C. meleagridis infection in the chicken poults was patchy and tended to be restricted to the lower part of the small intestine and in the ceca. The mouse gut was more intensely infected, and the infection was distributed throughout the small intestine and less distributed in the large intestine. The pyloric region of the stomach was free of infection, contrary to what is normally observed in mice infected with C. parvum (Akiyoshi et al., 2003). The intensity and distribution of infection and the mucosal alterations in piglets infected with C. meleagridis were milder than those seen with C. parvum but were similar to those of C. hominis (Akiyoshi et al., 2003). Overall, the rate of infectivity
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FIGURE 15.2 Electron micrographs of sections from a mouse (A to F and H) and a chicken (G) experimentally infected with Cryptosporidium meleagridis. (A) Early development of a trophozoite as it is being engulfed by the parasitophorous membrane. (B) Fully developed trophozoite. (C) First-generation schizogony with eight merozoites ready to be released into the intestinal lumen. (D) Second-generation schizogony with four merozoites that will develop into male or female sexual stages. (E) Dividing microgamy, showing 6 of 16 developing microgametes, the male sexual stage. (F) Fertilized macrogamete or zygote. (G) Fertilized macrogamete or zygote from an infected chicken, showing similar morphology and developmental stage to that of a zygote grown in a mouse (F). (H) Fully developed oocyst still within its parasitophorous vesicle. Bar, 1 µm. (Reproduced with permission from Akiyoshi, D.E., Dilo, J., Pearson, C., Chapman, S., Tumwine, J. and Tzipori, S. 2003. Characterization of Cryptosporidium meleagridis of human origin passaged through different host species. Infect. Immun. 71, 1828–1832.)
and virulence of C. meleagridis for the mammalian species was similar to that of C. parvum (Akiyoshi et al., 2003). Recently, a natural C. meleagridis infection was reported in a dog from the Czech Republic (Hajdusek et al., 2004). Although the dog was fed a commercial dry diet, the possible pseudoparasitic origin of C. meleagridis in dog feces cannot be ruled out. Whether dogs represent a natural host for C. meleagridis needs to be confirmed with more systematic characterization of cryptosporidiosis in dogs and by experimental infection studies. Oocysts resembling C. meleagridis in size have been detected in cases involving enteric cryptosporidiosis in quails, and it was suggested that an unnamed species of Cryptosporidium was responsible (Hoerr et al., 1986; Ritter et al., 1986; Guy et al., 1987; Lindsay and Blagburn, 1990). However, as C. meleagridis has been shown to have a wide host range, the species in question could be C. meleagridis. Genotyping the quail isolates is required to confirm this. Cryptosporidium meleagridis is also an emerging human pathogen and is the third most common Cryptosporidium parasite in humans (McLauchlin et al., 2000; Morgan et al., 2000a; Guyot et al., 2001; Pedraza-Diaz et al., 2001; Xiao et al., 2001; Yagita et al., 2001; Enemark et al., 2002; Gatei et al., 2002; Tiangtip and Jongwutiwes, 2002; Cama et al., 2003; Gatei et al., 2003; Leoni et al., 2003; Matos et al., 2004; Xiao et al., 2004; Coupe et al., 2005; Gatei et al., 2006a; Gatei et al., 2006b; Leoni et al., 2006a; Muthusamy et al., 2006). Cryptosporidium meleagridis in humans has been associated with gastrointestinal symptoms. In a recent study involving the molecular characterization of over 2000 human stool isolates of Cryptosporidium in the United Kingdom, 22 (~1%) were identified as C. meleagridis (PedrazaDiaz et al., 2001; Leoni et al., 2006a). In a study in Peru, C. meleagridis was more commonly identified in humans than C. parvum (Cama et al., 2003).
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403 Genetic Analysis
It has previously been suggested that C. meleagridis may be C. parvum (Champliaud et al., 1998; Sreter and Varga, 2000); however, molecular analysis of U.S., Australian, and European isolates at the small subunit ribosomal RNA (SSU rRNA), heat shock protein 70 (HSP70), Cryptosporidium oocyst wall protein (COWP), actin and glycoprotein 60 (GP60) loci, as well as the Cryptosporidium double-stranded RNA virus, have demonstrated the genetic and biological uniqueness of C. meleagridis (Xiao et al., 1999; Morgan et al., 2000b; Sreter et al., 2000; Sulaiman et al., 2000; Xiao et al., 2000; Morgan et al., 2001; Sulaiman et al., 2002; Xiao et al., 2002; Leoni et al., 2006b). In addition, when the morphology, host specificity, and organ location of C. meleagridis from a turkey in Hungary were compared with those of a C. parvum isolate, phenotypic differences were small but statistically significant (Sreter et al., 2000). Phylogenetic analyses at multiple loci have always placed C. meleagridis in a clade containing most mammalian Cryptosporidium parasites, and C. meleagridis is most related to the C. parvum, Cryptosporidium hominis, Cryptosporidium wrairi, the rabbit genotype, the mouse genotype, and the ferret genotype, all of which are parasites of the Euarchontoglires (primates, lagomorphs, and rodents) (Figure 15.3). Thus, C. meleagridis was possibly originally a mammalian Cryptosporidium parasite that subsequently became established in birds via ingestion of infected prey (Xiao et al., 2002). This is supported by the recent findings regarding the ability of C. meleagridis to infect a wide range of mammals (see Section D), which is certainly not the case with C. baileyi. Several subtypes of C. meleagridis have been described, based on multilocus analysis (Glaberman et al., 2001). Two subtypes of C. meleagridis were seen in the SSU rRNA gene, which differed from each other in the number of VspI restriction sites. Of the 11 human and bird isolates characterized, 6 subtypes were seen in the HSP70 and GP60 genes, indicating there is high genetic heterogeneity. Nevertheless, host isolation in C. meleagridis has not been seen thus far in C. meleagridis (Glaberman et al., 2001).
VII. Cryptosporidium baileyi Current, Upton and Haynes, 1986 A.
Biology
A second species of avian Cryptosporidium, originally isolated from commercial broiler chickens, was named C. baileyi based on its life cycle and morphologic features (Current et al., 1986) (Figure 15.4). The species was named in honor of the late W.S. Bailey, then President of Auburn University, for his pioneering work on the biology of Spirocerca lupi. The prepatent period was 3 days and the patent period lasted 20 and 10 days in birds inoculated at 2 days of age and at 1 or 6 months of age, respectively. Developmental stages, found in the microvillus region of enterocytes of the ileum and large intestine, were described, measured, and photographed and the time of their first appearance was noted. Heavy infection of the BF and cloaca did not result in clinical illness. Thin-walled oocysts were observed, but most were thick walled. Viable oocysts measured 6.3 × 5.2 µm and thus were much larger and more elongate than those of C. meleagridis (Table 15.1).
B.
Life Cycle
The life cycle of C. baileyi has been described in detail (Current et al., 1986). The prepatent period for C. baileyi was 3 days PI, and the patent period was 4 to 24 days for chickens inoculated at 2 days of age and 4 to 14 days for chickens inoculated at 1 and 6 months of age. During the first 3 days PI, most developmental stages of C. baileyi were found in the microvillous region of enterocytes of the ileum and large intestine. By day 4 PI, most parasites occurred in enterocytes of the cloaca and BF. Mature Type I meronts with 8 merozoites first appeared 12 h PI and measured 5.0 × 4.9 µm. Mature Type II meronts with 4 merozoites and a large granular residuum first appeared 48 h PI and measured 5.1 × 5.1 µm. Type III meronts with 8 short merozoites and a large homogeneous residuum first appeared 72 h PI and measured 5.2 × 5.1 µm. Microgamonts (4.0 × 4.0 µm) produced approximately 16 microgametes
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C. wrairi C. parvum C. hominis Ferret genotype 61
C. meleagridis Mouse genotype Skunk genotype
C. suis Fox genotype C. felis Cervid genotype
64
Marsupial genotype II
72
Opossum genotype I
98
Marsupial genotype I Muskrat genotype I
88
Deermouse genotype
Bear genotype C. canis C. varanii Pig genotype II
87
C. bovis
74 71
Duck genotype
78
Goose genotype I
94
Goose genotype II Snake spp. genotype 58 79
C. baileyi
94
Avian genotype I
88
99 Cryptosporidium sp. from ostrich Avian genotype II
Tortoise genotype 97 94
56
Avian genotype III Eurasian Woodcock genotype
100
C. serpentis 88 91 63
C. andersoni C. muris
92
Avian genotype IV 100
C. galli C. molnari-like genotype
Eimeria faurei Distance 0.1
FIGURE 15.3 Evolutionary relationships of avian Cryptosporidium isolates inferred by neighbor-joining analysis of Tamura–Nei distances calculated from pairwise comparison of SSU rRNA sequences. Percentage bootstrap support (> 60%) from 1000 pseudoreplicates is indicated at the left of the supported node.
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FIGURE 15.4 Scanning electron microscopy (SEM) pictures of a Cryptosporidium baileyi infection in the cloaca of a chicken. (Courtesy of W.L. Current, Lilly Research Laboratories, Indianapolis, IN.)
FIGURE 15.5 Scanning electron microscopy (SEM) pictures of Cryptosporidium baileyi life-cycle stages. (A) A single merozoite penetrating a host cell. (B) An unusual view of the basal portion of the feeder organelle after the parasite has become dislodged from the host cell. (C) Two Type I meronts. (D) A microgamete penetrating a macrogamete. (Courtesy of W.L. Current, Lilly Research Laboratories, Indianapolis, IN.)
that penetrated into macrogametes (4.7 × 4.7 µm). Macrogametes gave rise to two types of oocysts that sporulated within the host cells. Most were thick-walled oocysts (6.3 × 5.2 µm), the resistant forms that passed unaltered in the feces. Some were thin-walled oocysts whose wall (membrane) readily ruptured upon release from the host cell. Sporozoites from thin-walled oocysts were observed penetrating enterocytes in mucosal smears. The presence of thin-walled, autoinfective oocysts and the recycling of Type I meronts may explain why chickens develop heavy intestinal infections lasting up to 21 days. Some life-cycle stages are shown in Figure 15.5. A more recent study examined the ultrastructural features of sexual stages of C. baileyi in the respiratory tract of experimentally infected broiler chickens using transmission electron microscopy (Cheadle et al., 1999). Sexual stages of C. baileyi were seen attached to the tracheal epithelium and free in the tracheal lumen. These stages included intracellular Type III merozoite-like stages, microgamonts, microgametes, macrogamonts, thin-walled oocysts, and thick-walled oocysts. Thin-walled oocysts,
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microgamonts, and microgametes were seen less frequently than other sexual stages. Microgamonts, macrogamonts, and oocysts attached to the epithelium were all contained in a host cell membrane or within a parasitophorous vacuole. Thin-walled oocysts of C. baileyi were observed at the ultrastructural level in the respiratory tract of chickens (Cheadle et al., 1999).
C.
Pathogenesis
Natural C. baileyi infections have been reported in many anatomic sites in avian hosts, including the conjunctiva, nasopharynx, trachea, bronchi, air sacs, small intestine, large intestine, ceca, cloaca, BF, kidneys, and urinary tract (Lindsay and Blagburn, 1990). Following oral inoculation, the primary sites of development were the BF and cloaca (Lindsay et al., 1986). IT inoculation of oocysts into chickens resulted in extensive parasitization of the respiratory tract (Lindsay et al., 1986; Blagburn et al., 1987; Lindsay et al., 1987a). Conjunctival infections occurred in some birds when oocysts were placed directly on the conjunctival sac (Lindsay et al., 1987a). However, birds were more likely to develop infections in the BF and cloaca (Lindsay et al., 1987a). High morbidity and mortality have been associated with C. baileyi respiratory infections of birds, especially broiler chickens (Lindsay and Blagburn, 1990). Age-dependent resistance to C. baileyi infection was observed in chickens infected at 1 and 9 weeks of age (Sreter et al., 1995; Sreter et al., 2000). The prepatent period was significantly shorter and the patency was significantly longer in younger birds. Chickens infected at 1 week of age excreted thrice the number of oocysts excreted as those inoculated at 9 weeks of age. There was a good correlation between the length of the patent period and the total oocyst output of chickens (Sreter et al., 1995). Renal cryptosporidiosis was experimentally induced during a study to investigate the pathogenicity of C. baileyi in SPF chickens coinfected with Marek’s disease virus (MDV) (Abbassi et al., 1999). Cryptosporidium baileyi was administered orally at 4 days of age to chickens previously infected at hatching (day 0) with oncogenic MDV. Three control groups received MDV at hatching, C. baileyi on day 4, or a placebo consisting of distilled water. Renal cryptosporidiosis lesions were induced in the group coinfected with MDV and C. baileyi. The kidneys were markedly swollen and pale, with visible urate crystals in the ureters and surface tubules, and the presence of the parasite was confirmed by histologic examination of paraffin-embedded kidney sections. Histologic study also revealed subacute interstitial nephritis, acute ureteritis, and attachment of parasites on the epithelial cell surface of the ureters and collecting ducts, collecting tubules, and distal convoluted tubules. Various developmental stages of the parasite were present in the kidney sections (Abbassi et al., 1999).
D.
Host Range
Cryptosporidium baileyi is probably the most common avian Cryptosporidium species. It has been reported in a wide range of avian hosts, including black-headed gulls (Larus ridibundus), chickens, cormorants (Phalacrocorax spp.), a crane, a channel-billed toucan (Ramphastos vitellinus), an eastern golden-backed weaver (Ploceus jacksoni), turkeys, ducks, geese, cockatiels, a brown quail (Coturnix australis), a gray-bellied bulbul (Pycnonotus cyaniventris), an ostrich (Struthio camelus), a red-rumped cacique (Cacicus haemorrhous), a crested oropendola (Psarocolius decumanus), a red-crowned amazon (Amazona viridigenalis), a rose-ringed parakeet (Psittacula krameri), and a gray partridge (Perdix perdix) (Lindsay and Blagburn, 1990; Pavlásek, 1993; Ryan et al., 2003a; Abe and Iseki, 2004; Jellison et al., 2004; Kimura et al., 2004; Chvala et al., 2006). Experimental cross-transmission of C. baileyi to other birds, including Japanese quail, domestic ducks, geese, pheasants, a chukar partridge, and turkeys, were successful with the exception of bobwhite quail (Current et al., 1986; Lindsay et al., 1987a; Lindsay et al., 1989b; Lindsay and Blagburn, 1990; Cardozo et al., 2005). Limited life-cycle stages were observed in some turkey poults, and heavy infections developed only in the BF in 1-day and 2-day-old geese (Current et al., 1986). Sporozoites excysted in vitro and inoculated intranasally produced upper-respiratory infections similar to those reported for naturally infected broilers (Current et al., 1986). Mice and goats inoculated with C. baileyi oocysts, however, did not become infected (Current et al., 1986).
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TABLE 15.3 Genetic Similarities Between Avian Cryptosporidium Species and Genotypes at the SSU rRNA Locus Avian Avian Avian Avian Eurasian Goose C. C. C. Genotype Genotype Genotype Genotype Duck Woodcock Genotype baileyi meleagridis galli I II III IV Genotype Genotype I C. meleagridis C. galli Avian genotype I Avian genotype II Avian genotype III Avian genotype IV Duck genotype Eurasian Woodcock genotype Goose genotype I Goose genotype II
94.3 89.2 99.4
88.2 95.0
89.9
97.6
95.7
91.0
98.2
92.1
90.4
94.3
92.8
92.4
90.6
90.0
97.9
91.3
92.4
95.3
95.3
95.3
88.8
96.0
95.3
92.4
90.6
91.7
90.0
93.3
92.4
91.7
98.8
94.3
91.7
93.0
92.7
87.4
93.0
92.3
88.9
86.9
96.9
88.1
93.3
92.3
88.5
93.3
92.6
90.7
88.8
97.5
90.0
97.9
Note: Genetic similarities for Goose genotypes III and IV were not included because the partial sequence of the 18S rRNA available for comparison only overlapped for about 230 bp with the majority of isolates included here.
Cryptosporidium baileyi oocysts appear to be environmentally robust as studies have shown that chickens inoculated with C. baileyi oocysts that had been stored in 2.5% potassium dichromate at 4°C for 1 to 18 months developed patent infections, whereas oocysts stored for longer than this (up to 40 months) did not (Surl et al., 2003).
E.
Genetic Analysis
Cryptosporidium baileyi has been analyzed at a number of loci including the SSU rRNA, HSP70, COWP and actin loci and has been shown to be genetically distinct (Xiao et al., 1999; Sulaiman et al., 2000; Xiao et al., 2000; Morgan et al., 2001; Egyed et al., 2002; Sulaiman et al., 2002). Cryptosporidium baileyi is most closely related to avian genotype I and II (see Figure 15.13 and Table 15.3). The relationship between C. baileyi and the rest of the intestinal and gastric parasites is unclear as phylogenetic analysis at the SSU rRNA, actin, and COWP loci group C. baileyi with the intestinal parasites, but analysis at the HSP70 locus placed C. baileyi in a cluster that contained all gastric Cryptosporidium instead of the intestinal parasite cluster (Xiao et al., 2002).
VIII. Cryptosporidium galli Pavlásek, 1999 A.
Biology
A third species of avian Cryptosporidium was first described by Pavlásek (1999, 2001) in hens on the basis of biological differences. The parasite has recently been redescribed on the basis of both molecular and biological differences (Ryan et al., 2003b). Cryptosporidium galli oocysts are statistically different in size (p < 0.05) from C. baileyi (6.3 × 5.2 µm) and C. meleagridis oocysts (5.2 × 4.6 µm) and measure 8.25 × 6.3 (8.0–8.5 × 6.2–6.4) µm, with a length to width ratio of 1.3 (Pavlásek, 1999, 2001) (Figure 15.6 and 15.7 and Table 15.1). There is a spherical residual body, 3.6 to 4.0 µm in size, usually containing
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FIGURE 15.6 Nomarski Interference microscopy of Cryptosporidium galli oocysts. Bar, 10 µm. (Picture courtesy of Ivan Pavlásek, State Veterinary Institute Prague, Pathology and Parasitology Department, Prague, Czech Republic.)
FIGURE 15.7 Line drawing of Cryptosporidium galli. Bar = 5 µm.
three granules. Two of these granules, usually larger ones (approx. 1.6 µm in size), are positioned one against the other. The remaining granule is smaller (approx. 0.5–0.8 µm in size). The residual body is surrounded by four banana-shaped sporozoites, (approx. 12.8–14.4 × 0.8–1.0 µm in size) (Figure 15.8). Unlike other avian species, life-cycle stages of C. galli developed in epithelial cells of the proventriculus and not the respiratory tract or small and large intestines (Pavlásek, 1999, 2001). Cryptosporidium galli may also have been detected in birds when light and electron microscopy were used to characterize Cryptosporidium in the proventriculus of an Australian diamond firetail finch
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FIGURE 15.8 Nomarski interference microscopy of some Cryptosporidium galli life-cycle stages from proventriculum mucosal scrapings from a hen. (A) sporozoite, (B) cluster of trophozoites, (C) microgamont, (E) zygotes. (Pictures courtesy of Ivan Pavlásek, State Veterinary Institute Prague, Pathology and Parasitology Department, Prague, Czech Republic.)
FIGURE 15.9 H & E stained endogenous developmental stages of Cryptosporidium galli in the proventriculum of a naturally infected hen. (Picture courtesy of Ivan Pavlásek, State Veterinary Institute Prague, Pathology and Parasitology Department, Prague, Czech Republic.)
(Staganoplura bella) that died with acute diarrhea (Blagburn et al., 1990). A subsequent publication also identified a species of Cryptosporidium as infecting the proventriculus in finches and inadvertently proposed the name Cryptosporidium blagburni in Table 1 of the paper (Morgan et al., 2000b). However, Pavlásek (1999, 2001) had provided a detailed description of what appeared to be the same parasite and named it C. galli. More recent molecular analyses have revealed C. galli and C. blagburni to be the same species (Ryan et al., 2003a).
B.
Life Cycle
Little is known of the life cycle of C. galli. Endogenous developmental stages appear to be localized to glandular epithelial cells of the proventriculus. Life-cycle stages ranging from oocysts to trophozoites and macrogamonts have been observed (Figures 15.8 and 15.9).
C.
Pathogenesis
Cryptosporidium galli appears to be associated with clinical disease and high mortality (Blagburn et al., 1990; Pavlásek, 1999; Morgan et al., 2001; Pavlásek, 2001). Histopathology of infected finches
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demonstrated necrosis and hyperplasia of proventricular glandular epithelial cells associated with large numbers of Cryptosporidium oocysts attached to the surface of glandular epithelial cells (Morgan et al., 2001). Blagburn et al. (1990), gave a detailed histological description of a Cryptosporidium sp. that was probably C. galli infecting the proventriculus of an Australian diamond firetail finch. Infected proventricular glands had focal cuboidal metaplasia of epithelium, and developmental stages were observed within metaplastic segments of glands. Pale, eosinophilic, homogenous deposits occurred in the perivascular interstitial tissue at the base of the glands (Blagburn et al., 1990).
D.
Host Range
It appears that C. galli can infect a wide range of birds in different orders and families. Natural infections have been reported in finches, chickens, capercaille (Tetrao urogallus), pine grosbeak (Pinicola enucleator), turquoise parrots (Neophema pulchella), greater flamingo (Phoenicopterus ruber), red-cowled cardinals (Paroaria dominicana), and rhinoceros hornbills (Buceros rhinoceros) (Pavlásek, 1999, 2001; Ryan et al., 2003a; Ng et al., 2006). In addition, morphologically similar oocysts have been observed in a variety of exotic and wild birds including Phasianidae, Passeriformes, and Icteridae (Ryan et al., 2003b). A species that may have been C. galli has also been documented in the proventriculus of canaries (Tsai et al., 1983). Future studies are required to determine the extent of the host range for C. galli. Cross-transmission studies have shown that C. galli oocysts were infectious for 9-day-old but not 40day-old chickens (Pavlásek, 2001).
E.
Genetic Analysis
Distance, parsimony, and maximum likelihood analysis of three loci (SSU rRNA, HSP70, and actin) identified C. galli as a distinct species (Ryan et al., 2003b). The gastric location of C. galli in the host and its large size suggest that it is most closely related to the other gastric Cryptosporidium species, and this has been supported by molecular analysis (Figure 15.3). At the SSU rRNA locus, C. galli is most closely related to avian genotype IV and the gastric parasites Cryptosporidium muris, Cryptosporidium andersoni, and Cryptosporidium serpentis (Table 15.3), which is strongly supported by bootstrap analysis (Figure 15.3). At the SSU rRNA locus, C. galli isolates shared ~97.9% similarity with avian genotype IV, ~94.3% similarity with avian genotype III, ~96 % similarity with C. serpentis, and 95 to 96% similarity with C. andersoni and C. muris, respectively. The C. galli isolates shared only 89 to 93% similarity with other Cryptosporidium species outside of this cluster. At the HSP70 locus, C. galli isolates shared 94% similarity with C. serpentis and 93% similarity with C. muris. They shared 71 to 87% similarity with other Cryptosporidium species outside of this cluster. At the actin locus, C. galli isolates shared 96% similarity with C. serpentis and 93% similarity with C. muris and C. andersoni. They shared only 76 to 84% similarity with other Cryptosporidium species (Ryan et al., 2003b).
IX. Avian Genotypes A.
Avian Genotypes I–IV
Four distinct genotypes designated avian genotypes I–IV have been identified from various avian hosts (Meireles et al., 2006; Ng et al., 2006). Avian genotype I was identified in a red factor canary (Serinus canaria) (Table 15.4). Avian genotype II has been identified in an eclectus (Eclectus roratus), a galah (Eolophus roseicapillus), a cockatiel (Nymphicus hollandicus), and a Major Mitchell cockatoo (Cacatua leadbeateri) from Western Australia and from an ostrich from Brazil (Meireles et al., 2006). Sequence and distance analysis at the SSU rRNA gene locus indicated that although avian genotype I and avian genotype II were most closely related to C. baileyi, they were genetically distinct (~99.4 and 97.6% similarity, respectively, to C. baileyi) and exhibited ~98.2 % similarity to each other. At the actin gene locus, avian genotype I and avian genotype II exhibited only 95.7 and 88.3% similarity, respectively, to C. baileyi.
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TABLE 15.4 Avian Cryptosporidium Genotypes and their GenBank Accession Numbers for Various Loci Genotype
Host Species
Avian genotype I Avian genotype II
Red factor canary (Serinus canaria)
Avian genotype III
WA, Brazil
Eclectus (Eclectus roratus), WA, Brazil galah (Eolophus roseicapilla), cockatiel (Nymphicus hollandicus), Major Mitchell cockatoo (Cacatua leadbeateri, Ostrich (Struthio camelus) Galah (Eolophus WA, Brazil roseicapilla), cockatiel (Nymphicus hollandicus), sun conure (Aratinga solstitialis) Japanese whiteeye Czech (Zosterops japonica) Republic
Avian genotype IV Eurasian Eurasian woodcock woodcock (Scolopax rusticola) Duck Black duck (Anas genotype rubripes), Canada Geese (Branta canadensis) Goose Canada geese (Branta genotype canadensis) I Goose genotype II Goose genotype III Goose genotype IV
Geographic Origin
Canada geese (Branta canadensis)
SSU rRNA Locus Actin Locus DQ650339
HSP-70 Locus
DQ650346
Reference Ng et al., 2006
DQ650340, DQ650347, DQ650341, DQ650348, DQ002931 DQ002930
DQ002929 Meireles et al., 2006; Ng et al., 2006
DQ650342, DQ650349, DQ650343 DQ650350
NA
Ng et al., 2006
DQ650344
NA
NA
Ng et al., 2006
DQ650345
AY273773 Ryan et al., 2003; Ng et al., 2006 NA Morgan et al., 2001; Zhou et al., 2004; Jellison et al., 2004 NA Xiao et al., 2002; Zhou et al., 2004; Jellison et al. 2004
Czech AY273769 Republic Australia, AF316630, United States AY504514, AY324639 United States AY120912, AY504513, AY504516, AY324642 United States AY504512, AY504515, AY324643 United States AY324638
NA
NA
Zhou et al., 2004; Jellison et al., 2004
NA
NA
Jellison et al., 2004
AY324641
NA
NA
Jellison et al., 2004
Canada Geese (Branta canadensis)-isolate KLJ3b Canada geese (Branta United States canadensis )-isolate KLJ7
NA
AY120929
Note: NA = Not available.
Attempted transmission of avian genotype II oocysts to two groups of 2-day-old chickens did not result in infection as determined by histology, mucosal smears, and fecal screening until 4 weeks PI (Meireles et al., 2006). Avian genotype II has been reported to parasitize the cloacal epithelium and, to a lesser extent, the epithelium of the rectum and BF of ostriches (Santos et al., 2005). An earlier study reported a Cryptosporidium sp. in the feces of 14 of 165 (8.5%) ostriches imported into Canada (Gajadhar, 1994). The mean size of 40 oocysts measured was 4.6 × 4.0 µm (range 3.9–6.1 × 3.3–5.0 µm) with a shape index (length/width ratio) of 1.15 (range 1.00 to 1.38). In cross-transmission experiments, this Cryptosporidium sp. failed to infect suckling mice, chickens, turkeys, or quail (Coturnix coturnix japonica). Molecular data are not available for these ostrich isolates and, therefore, it is not possible to determine if the species described is the same as avian genotype II. Avian genotype III was identified in a galah, a cockatiel, and a sun conure (Aratinga solstitialis) from Western Australia and genetically formed a distinct group, which clustered with the Eurasian woodcock (Scolopax rusticola) genotype (see section B) at the SSU rRNA locus and received high bootstrap support
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(Figure 15.3). The avian genotype III exhibited 98.8% genetic similarity to the Eurasian woodcock genotype and 98.3% genetic similarity to C. serpentis, and shared 95.7 to 97.5% similarity with the other gastric parasites (C. galli, C. muris, and C. andersoni) at the SSU rRNA locus. At the actin gene locus, avian genotype III exhibited 98.5, 94.6, and 96.6% genetic similarity to the Eurasian woodcock, C. serpentis, and C. galli, respectively. Oocysts from avian genotype III measured 7.5 × 6.0 µm, which is larger than the mean oocyst size of C. serpentis (5.94 × 5.11 µm) but smaller than the oocyst size of the Eurasian woodcock-derived oocysts (8.5 × 6.4 µm). No clinical signs, such as diarrhea, dyspnea, coughing, or sneezing, were detected in the avian hosts for avian genotypes I to III. Avian genotype IV was identified in a Japanese whiteeye (Zosterops japonica) from the Czech Republic and exhibited ~97.9% similarity to its closest relative, C. galli, at the SSU rRNA locus. The Japanese whiteeye exhibited diarrhea and anorexia. Microscopic analysis detected only Cryptosporidium oocysts; no pathogenic bacteria were observed. Oocysts from avian genotype IV were similar in size to C. galli and measured 8.25 × 6.3 µm.
B.
Eurasian Woodcock Genotype
Another novel avian Cryptosporidium genotype was identified in a Eurasian woodcock (Scolopax rusticola) (isolate B2-1) from the Czech Republic (Ryan et al., 2003a). This bird had been obtained from the wild and transported to the Prague zoo. During the quarantine period, parasitological examination of the feces identified Cryptosporidium oocysts, which corresponded in size to the upper-limit dimensions of C. galli (i.e., 8.5 × 6.4 µm). However, the inner structure, particularly the size and shape of the rest body and granules, was different. During the week, the woodcock died and at autopsy, all endogenous developmental stages, including oocysts, were detected in the proventriculus only. At the SSU rRNA, actin and HSP70 loci (Ryan et al., 2003a; Ng et al., 2006) this genotype was shown to be genetically distinct and grouped most closely with avian genotype III and the gastric parasites (C. serpentis, C. muris, and C. andersoni) (Figure 15.3).
C.
Duck Genotype
A novel genotype was identified in a black duck (Anas rubripes) (Morgan et al., 2001), which appears to be most closely related to goose genotypes I and II at the SSU rRNA locus (96.9 to 97.5% similarity, respectively). Microscopic analysis of hematoxylin and eosin (H & E)-stained sections from the black duck indicated an enteric infection with numerous oocysts attached to the apical surface of the enterocytes of the small intestine, but clinical information on this bird was not available. The duck genotype has subsequently been identified in Canada geese (Branta canadensis) (Jellison et al., 2004; Zhou et al., 2004).
D.
Goose Genotypes I–IV
A recent study identified five Cryptosporidium sp. and genotypes from Canada geese collected from 13 sites in Ohio and Illinois; goose genotype I, goose genotype II, the duck genotype, C. parvum, and C. hominis (Zhou et al., 2004). Cryptosporidium goose genotypes I and II were the most common Cryptosporidium species in the fecal specimens studied; they had prevalence rates of 17.2 and 4.3%, respectively, were detected at 9 and 6 of the 13 study sites, respectively, and constituted 91.8% of the positive specimens. The high occurrence of these two Cryptosporidium species indicates that they are likely to be true parasites of Canada geese. Another related Cryptosporidium species, the duck genotype (Morgan et al., 2001), was found in one goose together with goose genotype I. It remains to be determined whether this parasite is infectious to Canada geese. The two other species, C. parvum and C. hominis, were only found in five geese, reaffirming the previous conclusion that oocysts of these two species were merely passing through the digestive tracts of foraging Canada geese without establishing infection (Graczyk et al., 1997). Phylogenetic and sequence analysis of the SSU rRNA gene has revealed that goose genotypes I and II and the duck genotype are closely related (Figure 15.3 and Table 15.3). These three avian parasites clustered together in a neighbor-joining tree (Figure 15.3). Goose genotypes I and II are
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closely related, forming a group within the cluster, with high bootstrap support. There also appears to be genetic variation within goose genotypes I and II as two subdivisions in both goose genotypes have been reported (Zhou et al., 2004). Another study identified five Cryptosporidium genotypes in Canada geese in the United States (Jellison et al., 2004). Three of the Cryptosporidium genotypes belonged to goose genotypes I (geese 1, 2, 3a, 6, and 8) and II (goose 9) and the duck genotype (goose 5), whereas the remaining two genotypes (geese 3b and 7) represented new Cryptosporidium genotypes: goose genotype III and IV, respectively.
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Hatkin, J., Giambrone, J.J. and Blagburn, B.L. 1993. Correlation of circulating antibody and cellular immunity with resistance against Cryptosporidium baileyi in broiler chickens. Avian Dis. 37, 800–804. Hoerr, F.J., Ranck, F.M., Jr. and Hastings, T.F. 1978. Respiratory cryptosporidiosis in turkeys. J. Am. Vet. Med. Assoc. 173, 1591–1593. Hoerr, F.J., Current, W.L. and Haynes, T.B. 1986. Fatal cryptosporidiosis in quail. Avian Dis. 30, 421–425. Hornok, S., Sreter, T., Bekesi, L., Szell, Z. and Varga, I. 1996. Attempts to immunize chickens with Cryptosporidium baileyi oocyst extract. J. Parasitol. 82, 650–652. Hornok, S., Heijmans, J.F., Bekesi, L., Peek, H.W., Dobos-Kovacs, M., Dren, C.N. and Varga, I. 1998. Interaction of chicken anaemia virus and Cryptosporidium baileyi in experimentally infected chickens. Vet. Parasitol. 76, 43–55. Hornok, S., Szell, Z., Shibalova, T.A. and Varga, I. 1999. Study on the course of Cryptosporidium baileyi infection in chickens treated with interleukin-1 or indomethacin. Acta Vet. Hung. 47, 207–216. Hornok, S., Szell, Z., Sreter, T., Kovacs, A. and Varga, I. 2000. Influence of in ovo administered Cryptosporidium baileyi oocyst extract on the course of homologous infection. Vet. Parasitol. 89, 313–319. Huang, K., Akiyoshi, D.E., Feng, X.C. and Tzipori, S. 2003. Development of patent infection in immunosuppressed C57BL/6 mice with a single Cryptosporidium meleagridis oocyst. J. Parasitol. 89, 620–622. Itakua, C., Goryo, M. and Umemura, T. 1984. Cryptosporidial infection in chickens. Avian Pathol. 13, 487. Jellison, K.L., Distel, D.L., Hemond, H.F. and Schauer, D.B. 2004. Phylogenetic analysis of the hypervariable region of the 18S rRNA gene of Cryptosporidium oocysts in feces of Canada geese (Branta canadensis): Evidence for five novel genotypes. Appl. Environ. Microbiol. 70, 452–458. Kimura, A., Suzuki, Y. and Matsui, T. 2004. Identification of the Cryptosporidium isolate from chickens in Japan by sequence analyses. J. Vet. Med. Sci. 66, 879–881. Latimer, K.S., Goodwin, M.A. and Davis, M.K. 1988. Rapid cytologic diagnosis of respiratory cryptosporidiosis in chickens. Avian Dis. 32, 826–830. Leoni, F., Gallimore, C.I., Green, J. and McLauchlin, J. 2003. 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. Leoni, F., Amar, C., Nichols, G., Pedraza-Diaz, S. and McLauchlin, J. 2006a. Genetic analysis of Cryptosponidium from 2414 humans with diarrhoea in England between 1985 and 2000. J. Med. Microbiol. 55, 703–707. Leoni, F., Gallimore, C.I., Green, J. and McLauchlin, J. 2006b. Characterisation of small double stranded RNA molecule in Cryptosporidium hominis, Cryptosporidium felis and Cryptosporidium meleagridis. Parasitol. Int. 55, 299–306. Levine, N.D., 1961. Protozoan Parasites of Domestic Animals and Man. Burgess Publishing Company, Minneapolis. Ley, D.H., Levy, M.G., Hunter, L., Corbett, W. and Barnes, H.J. 1988. Cryptosporidia positive rates of avian necropsy accessions determined by examination of auramine o-stained fecal smears. Avian Dis. 32, 108–113. Lindsay, D.S., Blagburn, B.L., Sundermann, C.A., Hoerr, F.J. and Ernest, J.A. 1986. Experimental Cryptosporidium infections in chickens: Oocyst structure and tissue specificity. Am. J. Vet. Res. 47, 876–879. Lindsay, D.S., Blagburn, B.L., Hoerr, F.J. and Giambrone, J.J. 1987a. Experimental Cryptosporidium baileyi infections in chickens and turkeys produced by ocular inoculation of oocysts. Avian Dis. 31, 355–357. Lindsay, D.S., Blagburn, B.L., Sundermann, C.A. and Ernest, J.A. 1987b. Chemoprophylaxis of cryptosporidiosis in chickens, using halofuginone, salinomycin, lasalocid, or monensin. Am. J. Vet. Res. 48, 354–355. Lindsay, D.S., Blagburn, B.L., Sundermann, C.A. and Giambrone, J.J. 1988a. Effect of broiler chicken age on susceptibility to experimentally induced Cryptosporidium baileyi infection. Am. J. Vet. Res. 49, 1412–1414. Lindsay, D.S., Sundermann, C.A. and Blagburn, B.L. 1988b. Cultivation of Cryptosporidium baileyi—Studies with cell-cultures, avian embryos, and pathogenicity of chicken embryo-passaged oocysts. J. Parasitol. 74, 288–293. Lindsay, D.S., Blagburn, B.L. and Sundermann, C.A. 1989a. Morphometric comparison of the oocysts of Cryptosporidium meleagridis and Cryptosporidium baileyi from birds. Proc. Helm. Soc. Wash. 56, 91–92. Lindsay, D.S., Blagburn, B.L., Sundermann, C.A. and Hoerr, F.J. 1989b. Experimental infections in domestic ducks with Cryptosporidium baileyi isolated from chickens. Avian Dis. 33, 69–73.
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Lindsay, D.S. and Blagburn, B.L., 1990. Cryptosporidiosis in Birds in Cryptosporidiosis in Man and Animals, Dubey, J.P., Speer, C.A. and Fayer, R., Eds. CRC Press, Boca Raton, FL, p. 133–148. Lindsay, D.S., Blagburn, B.L., Hoerr, F.J. and Smith, P.C. 1991. Cryptosporidiosis in zoo and pet birds. J. Protozool. 38, S180–S181. Mason, R.W. and Hartley, W.J. 1980. Respiratory cryptosporidiosis in a peacock chick. Avian Dis. 24, 771–776. Matos, O., Alves, M., Xiao, L.H., Cama, V. and Antunes, F. 2004. Cryptosporidium felis and C. meleagridis in persons with HIV, Portugal. Emerging Infect. Dis. 10, 2256–2257. McLauchlin, J., Amar, C., Pedraza-Diaz, S. and Nichols, G.L. 2000. 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. Meireles, M.V., Soares, R.M., dos Santos, M.M.A.B. and Gennari, S.M. 2006. Biological studies and molecular characterization of a Cryptosporidium isolate from ostriches (Struthio camelus). J. Parasitol. 92, 623–626. Morgan, U., Weber, R., Xiao, L.H., Sulaiman, I., Thompson, R.C.A., Ndiritu, W., Lal, A., Moore, A. and Deplazes, P. 2000a. Molecular characterization of Cryptosporidium isolates obtained from human immunodeficiency virus-infected individuals living in Switzerland, Kenya, and the United States. J. Clin. Microbiol. 38, 1180–1183. Morgan, U.M., Xiao, L., Limor, J., Gelis, S., Raidal, S.R., Fayer, R., Lal, A., Elliot, A. and Thompson, R.C.A. 2000b. Cryptosporidium meleagridis in an Indian ring-necked parrot (Psittacula krameri). Aust. Vet. J. 78, 182–183. Morgan, U.M., Monis, P.T., Xiao, L.H., Limor, J., Sulaiman, I., Raidal, S., O’Donoghue, P., Gasser, R., Murray, A., Fayer, R., Blagburn, B.L., Lal, A.A. and Thompson, R.C.A. 2001. Molecular and phylogenetic characterisation of Cryptosporidium from birds. Int. J. Parasitol. 31, 289–296. Murakami, S., Miyama, M., Ogawa, A., Shimada, J. and Nakane, T. 2002. Occurrence of conjunctivitis, sinusitis and upper region tracheitis in Japanese quail (Coturnix coturnix japonica), possibly caused by Mycoplasma gallisepticum accompanied by Cryptosporidium sp. infection. Avian Pathol. 31, 363–370. Muthusamy, D., Rao, S.S., Ramani, S., Monica, B., Banerjee, I., Abraham, O.C., Mathai, D.C., Primrose, B., Muliyil, J., Wanke, C.A., Ward, H.D. and Kang, G. 2006. Multilocus genotyping of Cryptosporidium sp. isolates from human immunodeficiency virus infected individuals in South India. J. Clin. Microbiol. 44, 632–634. Nakamura, K. and Abe, F. 1988. Respiratory (especially pulmonary) and urinary infections of Cryptosporidium in layer chickens. Avian Pathol. 17, 703–711. Ng, J., Pavlásek, I. and Ryan, U. 2006. Identification of novel Cryptosporidium genotypes from avian hosts. Appl. Environ. Microbiol. 72(12): 7548–7553. O’Donoghue, P.J., Tham, V.L., Desaram, W.G., Paull, K.L. and McDermott, S. 1987. Cryptosporidium infections in birds and mammals and attempted cross-transmission studies. Vet. Parasitol. 26, 1–11. O’Donoghue, P.J. 1995. Cryptosporidium and cryptosporidiosis in man and animals. Int. J. Parasitol. 25, 139–195. Pavlásek, I. 1993. Black headed gull (Larus ridibundus L), a new host of Cryptosporidium baileyi (Apicomplexa, Cryptosporidiidae). Vet. Med. (Praha) 38, 629–638. Pavlásek, I. 1999. Cryptosporidia: Biology, diagnosis, host spectrum, specificity, and the environment. Rem. Klin. Mikrobiol. 3: 290–301. Pavlásek, I. 2001. Findings of Cryptosporidia in the stomach of chickens and of exotic and wild birds. Veterinarstvi 51, 103–108. Pedraza-Diaz, S., Amar, C.F.L., McLauchlin, J., Nichols, G.L., Cotton, K.M., Godwin, P., Iversen, A.M., Milne, L., Mulla, J.R., Nye, K., Panigrahl, H., Venn, S.R., Wiggins, R., Williams, M. and Young, E.R. 2001. Cryptosporidium meleagridis from humans: Molecular analysis and description of affected patients. J. Infect. 42, 243–250. Proctor, S.J. and Kemp, R.L. 1974. Cryptosporidium anserinum sp. n. (Sporozoa) in a domestic goose Anser anser L., from Iowa. J. Protozool. 21, 664–666. Ranck, F.M. and Hoerr, F.J. 1987. Cryptosporidia in the respiratory tract of turkeys. Avian Dis. 31, 389–391. Randall, C.J. 1986. Renal and nasal cryptosporidiosis in a junglefowl (Gallus sonneratii). Vet. Rec. 119, 130–131. Ritter, G.D., Ley, D.H., Levy, M., Guy, J. and Barnes, H.J. 1986. Intestinal cryptosporidiosis and reovirus isolation from bobwhite quail (Colinus virginianus) with enteritis. Avian Dis. 30, 603–608.
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Ryan, U., Xiao, L.H., Read, C., Zhou, L., Lal, A.A. and Pavlásek, I. 2003a. Identification of novel Cryptosporidium genotypes from the Czech Republic. Appl. Environ. Microbiol. 69, 4302–4307. Ryan, U.M., Xiao, L., Read, C., Sulaiman, I.M., Monis, P., Lal, A.A., Fayer, R. and Pavlásek, I. 2003b. A redescription of Cryptosporidium galli Pavlasek, 1999 (Apicomplexa : Cryptosporidiidae) from birds. J. Parasitol. 89, 809–813. Santos, M.M., Piero, J.R. and Meireles, M.V. 2005. Cryptosporidium infection in ostriches (Struthio camelus) in Brazil: Clinical, morphological and molecular studies. Braz. J. Poult. Sci. 7, 109. Slavin, D. 1955. Cryptosporidium meleagridis (sp. nov.). J. Comp. Pathol. 65, 262–266. Sreter, T., Varga, I. and Bekesi, L. 1995. Age dependent resistance to Cryptosporidium baileyi infection in chickens. J. Parasitol. 81, 827–829. Sreter, T., Kovacs, G., Da Silva, A.J., Pieniazek, N.J., Szell, Z., Dobos-Kovacs, M., Marialigeti, K. and Varga, I. 2000. Morphologic, host specificity, and molecular characterization of a Hungarian Cryptosporidium meleagridis isolate. Appl. Environ. Microbiol. 66, 735–738. Sreter, T. and Varga, I. 2000. Cryptosporidiosis in birds—A review. Vet. Parasitol. 87, 261–279. Sreter, T., Szell, Z. and Varga, I. 2002. Anticryptosporidial prophylactic efficacy of enrofloxacin and paromomycin in chickens. J. Parasitol. 88, 209–211. Sulaiman, I.M., Morgan, U.M., Thompson, R.C.A., Lal, A.A. and Xiao, L.H. 2000. Phylogenetic relationships of Cryptosporidium parasites based on the 70-kilodalton heat shock protein (HSP70) gene. Appl. Environ. Microbiol. 66, 2385–2391. Sulaiman, I.M., Lal, A.A. and Xiao, L.H. 2002. Molecular phylogeny and evolutionary relationships of Cryptosporidium parasites at the actin locus. J. Parasitol. 88, 388–394. Sundermann, C.A., Lindsay, D.S. and Blagburn, B.L. 1987a. In vitro excystation of Cryptosporidium baileyi from chickens. J. Protozool. 34, 28–30. Sundermann, C.A., Lindsay, D.S. and Blagburn, B.L. 1987b. Evaluation of disinfectants for ability to kill avian Cryptosporidium oocysts. Companion Anim. Pract. 1, 36–39. Surl, C.G., Kim, S.M. and Kim, H.C. 2003. Viability of preserved Cryptosporidium baileyi oocysts. Korean J. Parasitol. 41, 197–201. Tacconi, G., Pedini, V., Gargiulo, A.M., Coletti, M. and Piergili-Fioretti, D. 2001. Retrospective ultramicroscopic investigation on naturally cryptosporidial infected commercial turkey poults. Avian Dis. 45, 688–695. Tarwid, J.N., Cawthorn, R.J. and Riddell, C. 1985. Cryptosporidiosis in the respiratory tract of turkeys in Saskatchewan. Avian Dis. 29, 528–532. Taylor, M.A., Catchpole, J., Norton, C.C. and Green, J.A. 1994. Variations in oocyst output associated with Cryptosporidium baileyi infections in chickens. Vet. Parasitol. 53, 7–14. Tham, V.L., Kniesberg, S. and Dixon, B.R. 1982. Cryptosporidiosis in quails. Avian Pathol. 11, 619. Tiangtip, R. and Jongwutiwes, S. 2002. Molecular analysis of Cryptosporidium species isolated from HIVinfected patients in Thailand. Trop. Med. Int. Health 7, 357–364. Trampel, D.W., Pepper, T.M. and Blagburn, B.L. 2000. Urinary tract cryptosporidiosis in commercial laying hens. Avian Dis. 44, 479–484. Tsai, S.S., Ho, L.F., Chang, C.F. and Chu, R.M. 1983. Cryptosporidiosis in domestic birds. Zhonghua Min Guo Wei Sheng Wu Ji Mian Yi Xue Za Zhi 16, 307–313. Tumova, E., Skrivan, M., Marounek, M., Pavlásek, I. and Ledvinka, Z. 2002. Performance and oocyst shedding in broiler chickens orally infected with Cryptosporidium baileyi and Cryptosporidium meleagridis. Avian Dis. 46, 203–207. Tyzzer, E. E. 1929. Coccidiosis in gallinaceous birds. Am. J. Hyg. 10, 269. Varga, I., Sreter, T. and Bekesi, L. 1995. Potentiation of ionophorous anticoccidials with Duokvin—Battery trials against Cryptosporidium baileyi in chickens. J. Parasitol. 81, 777–780. Wages, D.P. and Ficken, M.D. 1989. Cryptosporidiosis and turkey viral hepatitis in turkeys. Avian Dis. 33, 191–194. Whittington, R.J. and Wilson, J.M. 1985. Cryptosporidiosis of the respiratory tract in a pheasant. Aust. Vet. J. 62, 284–285. Wunderlin, E., Wild, P. and Eckert, J. 1997. Comparative reproduction of Cryptosporidium baileyi in embryonated eggs and in chickens. Parasitol. Res. 83, 712–715.
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Xiao, L., Morgan, U.M., Limor, J., Escalante, A., Arrowood, M., Shulaw, W., Thompson, R.C.A., Fayer, R. and Lal, A.A. 1999. Genetic diversity within Cryptosporidium parvum and related Cryptosporidium species. Appl. Environ. Microbiol. 65, 3386–3391. Xiao, L., Limor, J., Morgan, U.M., Sulaiman, I.M., Thompson, R.C.A. and Lal, A.A. 2000. Sequence differences in the diagnostic target region of the oocyst wall protein gene of Cryptosporidium parasites. Appl. Environ. Microbiol. 66, 5499–5502. Xiao, L., Bern, C., Limor, J., Sulaiman, I., Roberts, J., Checkley, W., Cabrera, L., Gilman, R.H. and Lal, A.A. 2001. Identification of 5 types of Cryptosporidium parasites in children in Lima, Peru. J. Infect. Dis. 183, 492–497. Xiao, L., Sulaiman, I.M., Ryan, U.M., Zhou, L., Atwill, E.R., Tischler, M.L., Zhang, X.C. and Fayer, R. 2002. Host adaptation and host-parasite co-evolution in Cryptosporidium: implications for taxonomy and public health. Int. J. Parasitol. 32, 1773–1785. Xiao, L., Fayer, R., Ryan, U. and Upton, S.J. 2004. Cryptosporidium taxonomy: recent advances and implications for public health. Clin. Microbiol. Rev. 17, 72–97. Yagita, K., Izumiyama, S., Tachibana, H., Masuda, G., Iseki, M., Furuya, K., Kameoka, Y., Kuroki, T., Itagaki, T. and Endo, T. 2001. Molecular characterization of Cryptosporidium isolates obtained from human and bovine infections in Japan. Parasitol. Res. 87, 950–955. Zhou, L., Kassa, H., Tischler, M.L. and Xiao, L. 2004. Host-adapted Cryptosporidium spp. in Canada geese (Branta canadensis). Appl. Environ. Microbiol. 70, 4211–4215.
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16 Zoo and Wild Mammals
Olga Matos
CONTENTS I. II.
Introduction .................................................................................................................................. 419 Cryptosporidiosis in Zoo and Wild Mammals ............................................................................ 420 A. Cryptosporidiosis in Wild Mammals .............................................................................. 420 1. Cryptosporidiosis in Wild Mammals: Cross-Transmission between Wild Mammals and Farm Animals ................................................................. 420 2. Ruminants ........................................................................................................ 420 3. Omnivores ........................................................................................................ 421 4. Carnivores ........................................................................................................ 421 5. Rodents............................................................................................................. 422 6. Lagomorphs...................................................................................................... 423 7. Insectivores....................................................................................................... 424 8. Bats................................................................................................................... 424 9. Marsupials ........................................................................................................ 424 10. Nonhuman Primates......................................................................................... 424 B. Zoo Mammals ................................................................................................................. 426 1. Zoo Animals..................................................................................................... 426 2. Research Animals............................................................................................. 426 3. Trade Animals .................................................................................................. 427 4. Control and Prevention .................................................................................... 427 III. Molecular Epidemiology of Cryptosporidiosis in Wild Mammals ............................................. 427 A. Genetic Diversity of Cryptosporidium in Wild Mammals............................................. 427 B. Species and Genotypes Detected in Wild Mammals ..................................................... 428 IV. Sources of Cryptosporidium Infection and Transmission Dynamics.......................................... 429 References........................................................................................................................... 430
I.
Introduction
Cryptosporidium infections have been reported in at least 155 mammalian species (O’Donoghue, 1995; Fayer et al., 1997; Fayer, 2004; Chapter 1, this book). The great majority of infections in mammals were reported in domestic animals of economic importance. Wildlife is considered an important source of infectious diseases transmissible to humans. Zoonoses from wildlife reservoirs constitute a major public health problem in all continents. The importance of such zoonoses is increasingly recognized, and the need for more attention in this area is being addressed. Wild animals seem to be involved in the epidemiology of most zoonoses and serve as major reservoirs for transmission of zoonotic agents to domestic animals and humans (Kruse et al., 2004). However, the involvement of wild animals in the epidemiology of cryptosporidiosis is less certain. Environmental pollution with human and domestic animal feces is recognized as a potential source for wildlife infections with zooanthropomorphic parasites such as Cryptosporidium and can put wildlife populations at risk (Appelbee et al., 2005). In addition, several wildlife species are parasitized by human-pathogenic 419
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Cryptosporidium species and have been frequently considered reservoirs of zoonotic disease and a source of environmental contamination (Xiao et al., 2000; Zhou et al., 2004; Jiang et al., 2005). Nevertheless, much of the research on Cryptosporidium in wild mammals has focused on cataloging species that are naturally susceptible to this parasite. Thus, the lack of morphological differences among species of Cryptosporidium found in animals has resulted in inaccurate information about host specificity and Cryptosporidium species and genotypes that were, in fact, involved in the infection. Only recently, the increased application of molecular methods in genotyping biological samples at informative loci has revealed that feral and captive wild mammals can harbor both host-adapted and zoonotic species and genotypes of Cryptosporidium, besides providing important insights into the taxonomy, host range, and zoonotic potential of this genus. Therefore, identification of Cryptosporidium species and genotypes and their susceptible hosts has contributed to the understanding of the transmission of cryptosporidiosis in wildlife and the public health significance of wildlife, and the formulation of measures to control cryptosporidiosis in humans and wild animals.
II. A. 1.
Cryptosporidiosis in Zoo and Wild Mammals Cryptosporidiosis in Wild Mammals Cryptosporidiosis in Wild Mammals: Cross-Transmission between Wild Mammals and Farm Animals
Infections by Cryptosporidium species have been documented in many wild mammals, and some researchers have suggested that these animals might constitute reservoirs and sources of infection for humans and domestic animals, even though few have fully assessed this issue. Cryptosporidium isolates from wild mice were found to infect calves (Klesius et al., 1986). In adult mammals, Cryptosporidium infection is mainly asymptomatic and the number of excreted oocysts is low and intermittent, even though it can persist for long periods (Casemore et al., 1997; Ramirez et al., 2004). It has been suggested, but largely remains undocumented, that free-ranging or pastured livestock could also constitute a source of infection for the wild animals with which they share a habitat. This was solely based on the theory that many of these wild animals, especially small mammals (rodents and insectivores) and cervids, are susceptible to C. parvum, which is frequently found in livestock (Olson et al., 2004). Recent studies, however, suggest that most of these animals are not infected with C. parvum, which is mainly a parasite of humans and preweaned calves (Zhou et al., 2004). The occurrence of cryptosporidiosis in wild animals has also been documented in nature reserves, some of which border rural areas where wild animals, humans, and domesticated cattle share habitats. For example, in a nature reserve in Tanzania, Mtambo et al. (1997) found cryptosporidiosis in 28% of 25 zebras (Equus zebra), 22% of 36 African buffalos (Syncerus caffer), and 27% of 26 wildebeest (Connochaetes gnou), whereas the infection rate in cattle near the park was 5.3% of 486 animals. Seasonal shifts of prevalence and oocyst shedding have been identified in studies involving wild animals in California, with infection more commonly seen in warm months (Atwill et al., 2004).
2.
Ruminants
Several studies have examined the prevalence of Cryptosporidium spp. in wild ruminants. Cryptosporidium species have been found in roe deer (Capreolus capreolus), fallow deer (Dama dama), sika deer (Cervus Nippon), Eld’s deer (Cervus eldithamin), barasingha deer (Cervus duvauceli), axis deer (Axis axis), mule deer (Odocoileus hermionus), white-tailed deer (Odocoileus virginianus), and caribou (Rangifer tarandus) (Korsholm and Henriksen, 1984; Heuschele et al., 1986; Rickard et al., 1999; Lourenço et al., 2000; Perz and Le Blancq, 2001; Siefker et al., 2002). Cryptosporidium parvum-like oocysts have also been detected in blackbuck (Antelope cervicapra), sable antelope (Hippotragus niger), scimitarhorned (Orix gazelle dammah) and fringe-eared (Orix gazelle callotys) orix, and in addax (Addax
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nasomaculatus) (Van Winkle, 1985; Heuschele et al., 1986). Cryptosporidium infection has also been reported in African buffalos (Syncerus caffer) and wildebeest (Connochaetes gnou) (Mtambo et al., 1997). In central Britain, C. parvum-like oocysts were found in feces of 6% of 16 fallow deer (D. dama) and 10% of 42 muntjac deer (Muntiacus reevesi). In each species, the mean number of oocysts/g of feces was 3000 (Sturdee et al., 1999). In Norway, fecal samples were collected from 1190 wild cervines and analyzed for cysts/oocysts of Giardia and Cryptosporidium (Hamnes et al., 2006). Cryptosporidium was found in 3.3% of 455 moose (Alces alces), 0.3% of 289 red deer (Cervus elaphus), and 6.2% of 291 roe deer (C. capreolus). Both Giardia and Cryptosporidium were found in samples from all geographical areas examined, indicating that these parasites are distributed among the cervine population in all parts of Norway, especially in moose and roe deer. In a natural park in Mafra, Portugal, Lourenço et al. (2000) detected oocysts of Cryptosporidium in 100% of 12 deer (Dama dama) studied, all in the range of 9 months to 4 years of age. In California, Cryptosporidium oocysts were detected in 15.9% of 82 asymptomatic, free-ranging adult cervines (Deng and Cliver, 1999). In white-tailed deer in several areas of Virginia and Mississippi, the prevalence of infection was 8.8 and 5%, respectively (Rickard et al., 1999). Perz and Le Blancq (2001) detected Cryptosporidium oocysts in fecal samples of 10.9% of 91 white-tailed deer inhabiting three areas in lower New York State. Cryptosporidium parvum-like oocysts have been detected in deer manure (Deng et al., 2000). Most studies of cryptosporidiosis in wild mammals report the prevalence of C. parvum-like oocysts, but the identity of the species or genotypes involved is not clear. In the Czech Republic, with the purpose of genotyping Cryptosporidium species in several animal and human hosts, C. parvum was identified in most deer isolates (Hajdusek et al., 2004). Cryptosporidium cervine genotype was identified in whitetailed deer in New York (Perz and Le Blancq, 2001). Cryptosporidium parvum of cervine origin has been experimentally transmitted to BALB/c and Porton mice (Tarazona et al., 1998). Cryptosporidium muris infections have been reported in gazelles and camels (Upton and Current, 1985; Iseki, 1986; Anderson, 1987; Pospischil et al., 1987; Fayer et al., 1991). This species has been experimentally transmitted to mice, rats, guinea pigs, rabbits, dogs, and cats (Iseki et al., 1989). The majority of the natural Cryptosporidium infections described in wild ruminants have been reported in zoo animals.
3.
Omnivores
In feral pigs that inhabit the edges of seeps, springs, ponds, and lakes in ten different areas along the coastal mountains of western California, the overall prevalence of Cryptosporidium was 5.4% of 221 animals (Atwill et al, 1997). Higher infection rates were found in animals younger than 8 months old or from high-density populations.
4.
Carnivores
The detection of Cryptosporidium in fecal samples of feral cats has been described in some publications (Mtambo et al., 1991; Cox et al., 2005). In a clinical and postmortem study of domestic and feral cats conducted in Glasgow, Scotland, 8.1% of 235 cats were infected with Cryptosporidium. More kittens than adults were infected (p < 0.01). There was no significant difference in the prevalence of Cryptosporidium infection between domestic and feral cats. Cryptosporidium oocysts were detected in fecal and mucosal impression smears, and endogenous developmental stages of the parasite were found in the microvillus region of enterocytes in 8 of 19 positive cats (Mtambo et al., 1991). In 100 feral cats and 76 domestic cats in North Carolina, Cryptosporidium oocysts were detected in the feces of 7% of feral cats and 6% of pet cats (Nutter et al., 2004). Specific anti-Cryptosporidium antibodies have been found in domestic and wild cats (Tzipori and Campbell, 1981; Mtambo et al., 1995). Sera from 258 healthy and sick domestic and feral cats, using an indirect immunofluorescence antibody test (IFA), were positive for IgG, IgM, and IgA antibodies in 74% (192/258), 32% (84/258), and 26% (67/258) of samples, respectively (Mtambo et al., 1995). Tzipori and Campbell (1981) also detected antibodies in 87% (20/23) of domestic cats. The detection of
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antibodies in such a high percentage of domestic and wild cats suggests widespread exposure to Cryptosporidium. Oocysts of Cryptosporidium species have been detected in feces of 9% of 23 foxes inhabiting mainland Britain (Sturdee et al., 1999) and in 7.9% of foxes and coyotes inhabiting wetlands adjacent to the Chesapeake Bay, Maryland (Zhou et al., 2004). Cryptosporidium infection has been reported in 15% of 26 badgers (Meles meles) (Sturdee et al., 1999) and in a striped skunk (Mephitis mephitis) in New York State (Perz and Le Blancq, 2001). Cryptosporidium found in a banded mongoose (Mungos mungo) represents a new genotype (Abe et al., 2004). Cryptosporidium infection has been reported in raccoons (Procyon lotor) (Carlson and Nielsen, 1982; Snyder, 1988; Martin and Zeidner, 1992; Zhou et al., 2004). Snyder found 13 of 100 wild raccoons infected, and 61% of the infected animals had a moderate-to-high parasite burden. Infection was localized in the small intestine of infected raccoons (Carlson and Nielsen, 1982; Martin and Zeidner, 1992). All infected raccoons were juveniles (Snyder, 1988; Martin and Zeidner, 1992). Cryptosporidium infection was found in 1 of 5 raccoons (Procyon lotor) from wildlife parks in New York State (Perz and Le Blancq, 2001) and in 2 of 51 raccoons in wetlands adjacent to the Chesapeake Bay (Zhou et al., 2004). Gastric cryptosporidial infections were never reported in raccoons. Based on molecular data, C. parvum was detected in raccoon dogs (Nyctereutes procyonoides viverrinus) (Matsubayashi et al., 2004; Abe et al., 2006).
5.
Rodents
Rodents are important in many ecosystems because they reproduce rapidly and can function as food sources for predators and as disease vectors. Their ubiquitous presence in rural environments and their sharing of habitat with farmed animals have led to epidemiological inquiries on the occurrence and identity of Cryptosporidium species in these animals. The first descriptions of the presence of Cryptosporidium oocysts in feces and its endogenous stages within tissues of mice were reported by Tyzzer in 1907 and 1910, followed by Hampton and Rosário in 1966. In Japan, the occurrence of Cryptosporidium oocysts in feces of brown rats (Rattus norvegicus) was reported in several studies. Iseki (1986) found a 10% prevalence of infection in 61, Miyaji et al. (1989) reported a 21% prevalence in 47, and Yamura et al. (1990) found a 2% prevalence in 48 brown rats. In the United Kingdom, Webster and Macdonald (1995) found C. parvum-like oocysts in 63% of 73 wild rats (R. norvegicus) trapped on nine rural farms around Oxfordshire and suggested that rats could represent a risk to human and livestock health. In three permanent populations of Norway rats in farmland in Warwickshire, United Kingdom, C. parvum-like oocysts were found in 24% of 438 rats (Quy et al., 1999). Cryptosporidium oocysts have also been detected in ship or black rats (Rattus rattus). Rates of 49% of 171 and 18% of 175 rats were reported in Japan (Miyaji et al., 1989; Yamura et al., 1990). In New Zealand (Chilvers et al. 1998), Cryptosporidium species was detected in 3 of 8 ship rats. Several studies reported C. parvum-like oocysts in feces of house mice (Mus domesticus). In the United States, 30% of 115 house mice were infected (Klesius et al., 1986). In the United Kingdom, C. parvum-like oocysts were detected in feces of 33% of 58 and 24% of 300 house mice (Chalmers et al., 1994, 1995). In New Zealand, oocysts were detected in feces of 11.8% of 17 house mice (Chilvers et al., 1998). Cryptosporidium was detected in 15% of 39 yellow-necked mice (Apodemus flavicollis) and in 20% of 275 bank voles (Clethrionomys glareolus) trapped in the District of Mazury Lake, Poland (Siski et al., 1993). Later, in the same area, Cryptosporidium oocysts were detected in 23% of 102 C. glareolus, ´ 24% of 70 Apodemus sp., and in 1 Apodemus agrarius, and 4 Microtis arvalis (Sinski et al., 1998). In Finland, cryptosporidial infection was detected in 2% of 41 bank voles and 1% of 131 field voles (Microtus agrestis) (Laakkonen et al., 1994). In various sites of the Province of Cataluna, Spain, infection with C. parvum-like oocysts was detected in 35.2% of 278 Apodemus sylvaticus, 27.3% of 22 Mus spretus, 20.4% of 49 C. glareolus, and in 1 R. rattus and 1 A. flavicollis (Torres et al, 2000). In Poland,
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Bajer et al. (2002) studied the prevalence of Cryptosporidium spp. in three species of wild rodents (A. flavicollis, C. glareolus, and M. arvalis) from forests and abandoned agricultural fields during 1997–1999, and found an overall prevalence of 62% (582/942). At an agricultural site in the United Kingdom, C. parvum-like oocysts were detected in 19% of 595 wild rodents (M. domesticus, A. sylvaticus, and C. glareolus) (Chalmers et al., 1997). All these rodents were asymptomatic. A longitudinal survey of Cryptosporidium in livestock and small wild mammals conducted over 6 years (1992–1997) on a farm in the same country found the parasite to be endemic and persistently present in all mammalian categories (Sturdee et al., 2003). Through a morphometric analysis of the detected oocysts, these authors found C. parvum-like oocysts in the feces of 32.8% of the rodents living in and around farm buildings and in 29.9% of the small wild mammals (mainly rodents) living in areas of pasture. The highest Cryptosporidium prevalence was reported in the autumn. In the United States, Cryptosporidium species were detected in 1 of 2 white-footed mice (Peromyscus leucopus) (Perz and LeBlanc, 2001). Cryptosporidium has been found in 5 of 9 muskrats (Ondatra ´ zibethicus) by conventional microscopy (Sinski et al., 1998). It was detected by molecular methods in 100% of 6 muskrats (Perz and Le Blancq, 2001) and in 11.8% of 237 muskrats (Zhou et al., 2004). Cryptosporidial infection has also been detected in 11% of 19 European beavers (Castor fiber) (Siski et al., 1998) and 3% of 62 North American beavers (Castor canadensis) (Fayer et al., 2006). Cryptosporidium stages were reported in the cecal epithelium of a young gray squirrel (Sciurus carolinensis) (Sundberg et al., 1982). It was reported in naturally infected asymptomatic Siberian chipmunks (Tamias sibiricus) from the Peoples Republic of China (Matsui et al., 2000). Oocysts morphologically similar to C. parvum were inoculated into SCID mice and developed in the lower jejunum and ileum. No parasites were detected in the stomach. Chipmunks (Tamias striatus) from parks in New York State were also found infected with Cryptosporidium (Perz and Le Blancq, 2001). In Kern County, CA, using molecular methods, Cryptosporidium was found in 16% of 309 ground squirrels (Spermophilus beecheyi) from 17 geographic locations (Atwill et all, 2001). Squirrels were shedding an average of 53,875 oocysts/g of feces. Male squirrels had higher prevalence and intensity of shedding than female squirrels. It was postulated that the higher intensities of shedding by males might increase dissemination and genotypic mixing of Cryptosporidium because of their proclivity to disperse to nonnatal colonies. In a subsequent study at the same location, 12% of 853 squirrels were shedding Cryptosporidium oocysts (Atwill et al., 2004). Shedding was higher in summer, and juveniles, particularly males, were about twice as likely as adults to be infected and shed higher numbers of oocysts. Cryptosporidium parvum-like oocysts were also detected in squirrels in the Italian Alps (Bertolino et al., 2003). Young squirrels were more frequently infected than adults. Cryptosporidium muris oocysts have been detected in rodents less frequently than C. parvum-like oocysts. Cryptosporidium muris-like oocysts were detected in 26% of 58, 13% of 300, and 10% of 242 naturally infected house mice (Chalmers et al., 1994, 1995, 1997). This species has also been detected in 5% of 230 wood mice (Chalmers et al., 1997), and in 2% of 123 and 43% of 114 bank voles (Chalmers et al., 1997; Bull et al., 1998). In Japan, naturally infected brown rats were reported to be shedding C. muris-like oocysts (Iseki, 1986). In Spain, C. muris-like oocysts were found in 3.9% of 278 A. sylvaticus and in 4% of 49 C. glareolus (Torres et al., 2000). Mixed infections with C. parvum and C. muris-like oocysts were detected in 5.8% of 278 A. sylvaticus, 4.5% of 22 M. spretus, and 4% of 49 C. glareolus (Torres et al., 2000). However, in an extensive study in the United Kingdom, C. muris was rarely found in small wild mammals (mainly rodents) (Sturdee et al., 2003). In the Czech Republic, oocysts of Cryptosporidium, morphologically similar to C. muris, were found in naturally infected Siberian chipmunks (Eutamias sibiricus). BALB/c mice, experimentally inoculated with these, began shedding oocysts 14 to 35 days post infection; and developmental stages of C. muris were found in the glandular stomach. Clinical signs were absent in both chipmunks and experimentally infected mice (Hurkóva et al., 2003).
6.
Lagomorphs
The first descriptions of Cryptosporidium in feces of rabbits were reported by Tyzzer in 1912. Developmental stages were described in 1979 in the intestinal tract of an asymptomatic adult female rabbit
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(Oryctolagus cuniculus). The organism was morphologically and developmentally indistinguishable from C. parvum. At that time it was considered to be host specific, and it was named C. cuniculus (Inman and Takeuchi, 1979). In the same year, there was another report of C. cuniculus identified by light and electron microscopy in two apparently healthy rabbits (Rehg et al., 1979). The subsequent transmission of C. parvum from calves to rabbits led to the suggestion that C. cuniculus was synonymous with C. parvum (Fayer and Ungar, 1986). Since then, Cryptosporidium infections have been reported in commercial (Peeters et al., 1986), farmed (Ryan et al., 2003), laboratory (Iseki et al., 1989; Mosier et al., 1997), and wild rabbits (DaSilva et al., 1992). Cryptosporidium has also been detected in a wild cottontail rabbit (Sylvilagus floridanus) (Ryan et al., 1986). In Norfolk, United Kingdom, C. parvum-like oocysts were detected in 7% (2/28) of live-trapped rabbits (O. cuniculus), with an estimate of 3000 oocysts/g of feces (Sturdee et al., 1999).
7.
Insectivores
Cryptosporidium oocysts were detected in feces of 31% of 16 naturally infected common shrews (Sorex ´ araneus) in the District of Mazury Lake, Poland (Sinski et al., 1993). Cryptosporidial infection in a hedgehog was reported for the first time by Graczyk et al. (1998). In 1999, Sturdee et al. detected C. parvum-like oocysts in 1 of 4 hedgehogs (Erinaceus europaeus), in 35% of 20 common shrews (S. araneus), and in 1 of 10 pygmy shrews (Sorex minutus)—species indigenous or naturalized to mainland Britain (Sturdee et al., 1999). In the Province of Cataluna, Spain, Cryptosporidium oocysts were detected in 14.8% of 88 Crocidura russula shrews (Torres et al., 2000).
8.
Bats
Cryptosporidium infections were also reported in two species of bats (Eptesicus fuscus and Myotus adversus) (Dubey et al., 1998; Morgan et al., 1999).
9.
Marsupials
Developmental stages of Cryptosporidium were observed in histological sections of the small intestine from male brown antechinus (Antechinus stuartii) and from five koalas (Phascolarctos cinereus) from a wild park in Australia, and in histological sections of the intestine of a hand-reared juvenile Tasmanian pademelon (Thylogale billardierii) (Barker et al., 1978; O’Donoghue, 1995). Oocysts similar to those of C. parvum were detected in feces of two southern brown bandicoots (Isoodon obesulus), a ratlike marsupial, and a hand-reared orphan juvenile red kangaroo (Macropus rufus), in South Australia (O’Donoghue, 1995). Cryptosporidium oocysts were isolated from feces collected from eastern grey kangaroos (Macropus giganteus) inhabiting an Australian water catchment and characterized by molecular methods (Power et al., 2004). The prevalence of Cryptosporidium in this host population was estimated to range from 0.32 to 28.5%, with peak infections during the autumn months. Oocyst shedding intensity ranged from less than 20 oocysts/g feces to 2.0 × 106 oocysts/g feces, and shedding did not appear to be associated with diarrhea (Power et al., 2005). Cryptosporidiosis appears to be a mild infection in opossums (Didelphis virginiana). Five nursing opossums experimentally inoculated with 5 × 106 C. parvum oocysts of calf origin were susceptible to the infection (Lindsay et al., 1988). Developmental stages of C. parvum were observed in the ileum, cecum, and colon of these opossums. Two of three uninoculated pouch mates acquired the infection, confirmed by the examination of feces and tissue sections. Seven of the C. parvum infected opossums had mild diarrhea, although none died as a result of the infection.
10.
Nonhuman Primates
Numerous natural and experimentally induced infections have been reported in nonhuman primates. Cryptosporidium infection with oocysts undistinguishable from C. parvum have been described in squirrel monkeys (Saimiri sciureus), red-ruffed lemurs (Varecia variegate rubra), brown lemurs (Lemur macuco-mayottensis), marmosets (Callithrix jacchus, Saguinus Oedipus), macaques (M. mulatta,
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Macaca nemestrina, Macaca fascicularis, Macaca fuscata), spider monkeys (Aetles belzebuth), mangabeys (Cercocebus torquatus lunulatus), green monkeys (Cercopithecus aethiops, Cercopithecus campbelli, Cercopithecus talapoin), patas monkeys (Erethrocebus patas), orangutan (Pongo pygmaeus), baboons (Papio anubis) (Kovatch and White, 1972; Heuschele et al., 1986; Riggs, 1990; Miller et al., 1990; Gómez et al., 1992; O’Donoghue, 1995; Kalishman et al., 1996; Fayer et al., 1997; Hope et al., 2004), and gorillas (Gorilla gorilla beringei) (Nizeyi et al., 1999; Graczyk et al., 2001). Cryptosporidium infection has been identified in 11 of 100 free-ranging mountain gorillas (G. gorilla beringei) in Uganda (Nizeyi et al., 1999) and in the free-ranging mountain gorillas of the Bwindi Impenetrable National Park, Uganda (Graczyk et al., 2001). Molecular characterization of two Cryptosporidium isolates originating from two human-habituated gorilla groups and two isolates from nonhabituated gorillas (G. gorilla beringei) yielded positive identification of C. parvum (Graczyk et al., 2001). Because some populations of free-ranging mountain gorillas have become habituated to humans, and as C. parvum is cross-transmissible between humans and animals, the authors concluded that C. parvum infections can be propagated in the habitats of human-habituated, free-ranging gorillas through both zoonotic and anthroponotic transmission cycles (Graczyk et al., 2001). After this report, Nizeyi et al. conducted a study to determine whether C. parvum infections were present in people sharing habitats with free-ranging mountain gorillas in the same National Park and, if so, to identify the genotype of Cryptosporidium causing infections in humans. Cryptosporidium infection was found in 21% of the park staff members who had frequent contact with gorillas in comparison with 3% of prevalence in the local community, indicating the existence of a zoonotic transmission cycle of this pathogen (Nizeyi et al., 2002a). The same group expanded the study to cattle grazing in the vicinity of the same national park. The prevalence of cryptosporidiosis and giardiasis was 38 and 12%, respectively, with 10% concomitant infections. Shedding intensity varied from 130 to 450 oocysts/g (mean of 215 oocysts/g) and from 110 to 270 cysts/g (mean of 156 cysts/g), respectively. Significantly, more preweaned than postweaned cattle were infected with either parasite, and the preweaned cattle shed significantly higher numbers of either parasite than the postweaned cattle. Mathematical modeling indicated that the maximum prevalence of asymptomatic infections reached approximately 80% for cryptosporidiosis and 35% for giardiasis in the sampled cattle. Because C. parvum recovered from cattle can infect people and gorillas, these authors concluded that cattle that graze within the Bwindi Park were a source of C. parvum infection for gorillas (Nizeyi et al., 2002). At four localities in the Rift Valley in Ethiopia, Cryptosporidium oocysts were detected in 11.9% of 59 baboons (Papio. anubis) and 29.3% of vervets (Cercopithecus aethiops) (Legesse and Erko, 2004). These primates use the same water sources as humans and range freely in human habitats, suggesting that (without molecular or epidemiological confirmation) they might be a source of oocysts infectious for humans. In Sri Lanka, Ekanayake et al. (2006) studied the prevalence of enteric parasites among nonhuman primates that inhabit the natural forests. A coprologic survey of 125 monkeys (89 toque macaques, 21 gray langurs, and 15 purple-faced langurs) found Cryptosporidium in all three species, especially among monkeys using areas and water heavily contaminated by humans and livestock. Most macaques (96%) shedding Cryptosporidium oocysts were coinfected with other protozoans and helminths (e.g., Enterobius and Strongyloides). In summary, the aforementioned reports suggest that transmission of Cryptosporidium might occur not only among primates in the wild but between them and humans, and, possibly, domesticated livestock. These findings could have important implications for public health as well as wildlife conservation management. Cryptosporidium rhesi, the name given to an isolate from a rhesus monkey (Macaca mulatta) (Levine, 1980), morphologically indistinguishable from C. parvum, is not considered a valid species (Riggs, 1990; Fayer et al., 1997; Chapter 1, this book).
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B.
Zoo Mammals
1.
Zoo Animals
The epidemiological studies carried out on cryptosporidiosis in nondomesticated mammals are very rare, and many times they are based on relatively small samples of groups in captivity in zoological parks. The majority of the natural infections described in wild mammals have been reported in zoo animals. The detection of Cryptosporidium has been described in various species of exotic ruminants and primates with diarrhea in captivity. At the San Diego Wild Animal Park, CA, Heuschele et al. (1986) detected cryptosporidial infections in 52.5% of 40 exotic ruminants with diarrhea, and in 2 primates with gastroenteritis. In the zoo of Barcelona, Spain, oocysts of Cryptosporidium were found in fecal samples of various species of ruminants (Giraffidae, Bovidae, Cervidae, Camelidae, Rinocerotidae, and Elefantidae) and primates (Lemuridae, Pongidae, Cercopithecidae, and Hapalidae) (Gómez et al., 1992, 1996, 2000). The prevalence of Cryptosporidium in ruminants increased from 17.6% of 34 in 1996 to 53.2% of 62 in 2000. In addition, 27.6% of the primates studied in 1996 were infected with Cryptosporidium, which increased to 44.4% in 2000. Clinical signs associated with Cryptosporidium in these animals varied from absence of clinical signs to severe diarrhea, which in some cases led to death. Oocysts were morphologically similar to C. parvum, although molecular characterization was not performed. In the zoo of Pozna, Poland, of 66 animals examined, oocysts were detected in feces of a lesser slow loris (Nycticebus pygmaeus), a Japanese macaque (M. fuscata), an Indian elephant (Elephas maximus), a white rhinoceros (Ceratotherium simum), an Eld’s deer (C. eldi), and a Thorold’s deer (Cervus albirostris) (Majewska et al., 1997). In most parasitized animals, the intensity of infection was light, and none of the animals had any apparent symptoms of cryptosporidiosis. In the zoo of Lisbon, Portugal, of 34 species of ruminants, 4% of 196 animals had cryptosporidiosis (Delgado et al., 2003), and molecular characterization identified the isolates as C. parvum (Alves et al., 2001, 2003). The same group expanded the investigation to a larger sample size and found Cryptosporidium oocysts in a black wildebeest (Connochaetes gnou) and a Prairie bison (Bison bison bison) (Alves et al., 2005). All animals were asymptomatic. Cryptosporidium muris was reported in mountain gazelles (Gazella cuvieri) from the zoo of Munich, Germany (Popischil et al., 1987). Cryptosporidiosis was reported in an adult female Bactrian camel (Camelus bactrianus) housed at the National Zoological Park in Washington, D.C. (Fayer et al., 1991). The parasite was experimentally established in mice, with oocysts morphologically identical to C. muris. Subsequent molecular characterization confirmed the species identification (Xiao et al., 1999; Morgan et al., 2000). The environmental temperature and humidity, the physical features of the facilities, the vicinity of the animals, and the captivity-induced stress may lead to immunosuppression (Estes et al., 1992), which may contribute to transmission of enteropathogens in zoological parks. Factors influencing the transmission of Cryptosporidium in primates and herbivores housed at the Barcelona Zoo were analyzed (Gracenea et al., 2002). The relationship of continuous and discontinuous oocyst shedding to animalhousing conditions and abiotic factors (seasonality, humidity, and temperature) were examined to determine the epizootiology of the cryptosporidiosis. After 36 fecal samples from each of 11 primates and 22 herbivores were examined over the period of a year, it was concluded that transmission primarily resulted from chronic infection in some animals serving as a source of successive reinfection for other animals. The environmental temperature and humidity (seasonality), the physical features of the facilities, the vicinity of the animals, and the physiological status induced by captivity contributed to cryptosporidiosis transmission.
2.
Research Animals
Animals kept in captivity for research purposes may be more prone to infection with Cryptosporidium because of the high density of animals and stressful living conditions. A case of spontaneous cryptosporidiosis was reported in an adult female white-tailed deer (O. virginianus) maintained in captivity for research at the University in Georgia (Fayer et al., 1996).
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In a captive primate colony at the Wisconsin Regional Primate Research Center, Cryptosporidium oocysts were detected in 4% of 25 marmosets older than 1 year and in 16% of 25 animals less than 1 year. All animals appeared healthy. These results suggest that captive marmosets of all ages are potential carriers of Cryptosporidium, although young animals seem to be more susceptible to infection (Kalishman et al., 1996). Cryptosporidium muris-like parasites were detected in gastric tissues from 15 cynomolgus monkeys (Macaca fascicularis) in a toxicity study of immunosuppressive drugs (Dubey et al., 2002). In captive lemurs (Propithecus verreauxi coquereli) in the Duke University Primate Center, North Carolina, sequence and phylogenetic analysis of oocysts from feces indicated that these primates were infected with the cervine genotype of Cryptosporidium (Da Silva et al., 2003). In Australia, C. muris infection was detected in 39.2% of 28 bilbies (Macrotis lagotis) housed at a captive breeding colony (Warren et al., 2003). A case of fatal cryptosporidiosis in a juvenile captive African hedgehog (Ateletrix albiventris) was confined to the ileum, jejunum, and colon infection, and was extremely severe in the lower jejunum, where over 75% of the epithelial cells harbored the pathogen (Graczyk et al., 1998).
3.
Trade Animals
Animals kept in captivity for trade purpose are prone to infections with Cryptosporidium spp. In recently captured or imported animals, cryptosporidial infections are frequently observed, which is probably related to the stress of the capture, transport, and temperature and food changes. In a study performed in the Czech Republic, C. muris was detected in Siberian chipmunks (Eutamias sibiricus) imported from Southeast Asia by a pet trader (Hurková et al., 2003). The Cryptosporidium ferret genotype was detected in fecal samples of ferrets exhibited at a pet shop in the city of Kanazawa, Japan (Abe and Iseki, 2003).
4.
Control and Prevention
Preventative measures are the most basic aspect of the medical care of captive wildlife (Miller, 1999), and parasitological surveys can help monitoring the diseases. Measures for controlling outbreaks of bovine cryptosporidiosis may be applicable to zoo animals (Harp and Goff, 1998). Keepers should thoroughly clean (or use separate) boots, protective clothing, and utensils when moving from one enclosure to another. Vehicle tires should be cleaned before leaving an enclosure to prevent the dissemination of organisms. Zoological parks are public facilities visited annually by multitudes of people. Visitors should be encouraged not to touch the animals or surfaces in contact with animals and to wash their hands before eating. It is of public health importance to monitor the prevalence of the infection in caretakers and the resident population, because the presence of animal carriers could facilitate zoonotic transmission.
III. Molecular Epidemiology of Cryptosporidiosis in Wild Mammals A.
Genetic Diversity of Cryptosporidium in Wild Mammals
Molecular studies have shown an extensive genetic diversity in Cryptosporidium species infecting mammals (humans, and domesticated and wild animals), reptiles, and birds. Based on multiple parameters, which include genetic characterization, morphometric studies, host specificity, and localization in the host, 10 Cryptosporidium species have been established in mammals—Cryptosporidium andersoni (bovines), Cryptosporidium hominis (humans), Cryptosporidium parvum (ruminants, humans), Cryptosporidium canis (canids, humans), Cryptosporidium felis (felids, humans), Cryptosporidium wrairi (guinea pig), Cryptosporidium suis (pigs, humans), Cryptosporidium muris (rodents, humans), Cryptosporidium meleagridis (birds, humans), and Cryptosporidium bovis (ruminants) (Xiao et al., 2004; Ryan et al., 2004; Fayer et al., 2005). In addition to these species, over 30 host-adapted genotypes have been described from mammals such as the monkey, rabbit, horse, ferret, mouse, skunk, opossum I and II, marsupial I and II, muskrat I and II, cervine, fox, squirrel A, B, and C, deer mouse, bear, wildebeest,
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C. canis coyote and C. canis fox, pig II, deer, deer-like, mongoose, and several squirrel genotypes (Morgan et al., 1999a; Xiao and Ryan 2004; Xiao et al., 2004, Alves et al, 2005). The demonstration of these species and genotypes of Cryptosporidium, most of them largely host specific, suggests that multiple routes of transmission are involved in the transmission in different host species.
B.
Species and Genotypes Detected in Wild Mammals
The zoonotic transmission of cryptosporidiosis to humans can result from direct contact with infected animals (Juranek, 1995; Casemore et al., 1997). Because C. parvum is a species normally, but not exclusively, associated with zoonotic transmission and appears to have a broad host range, it was generally believed that most Cryptosporidium infections in wild mammals, with oocysts morphometrically identical to C. parvum, were due to C. parvum. Based on this assumption, wild mammals were thought to represent an important zoonotic reservoir for human cryptosporidiosis. However, application of molecular methods to genotype samples at informative loci has revealed that feral and captive wildlife can harbor both hostadapted and zoonotic species and genotypes of Cryptosporidium. The only species of Cryptosporidium detected and confirmed by molecular methods in wild mammals are C. parvum, C. muris, and C. canis. Cryptosporidium parvum has been isolated from only a few wild mice, deer, raccoon dog, and gorillas (Morgan et al., 1999; Deng and Cliver, 1999; Perz and Le Blancq, 2001; Graczyk et al., 2001; Matsubayashi et al., 2004; Xiao et al., 2004; Xiao and Ryan, 2004; Alves et al., 2006; Abe et al., 2006). In all these instances, infections were asymptomatic. In comparison, C. muris infections have been essentially detected by morphometric analysis of oocysts. Applying both parasitological and molecular methods, C. muris was detected and confirmed in Siberian chipmunks imported from Southeast Asia by a pet trader (Hurková et al., 2003). In Australia, C. muris was detected in bilbies (Macrotis lagotis) by molecular methods (Warren et al., 2003). On a farm in Japan, C. muris type oocysts were detected in 21 of 516 beef cattle and in 2 of 25 Japanese field mice (Apodemus speciosus) on the same farm. Gene analysis suggested that the oocysts isolated from the wild mice were C. andersoni and C. muris (Nakai et al., 2004). Later, the same group inoculated the oocysts isolated from the mice into large Japanese field mice and SCID mice, and developing stages were found in the stomach epithelium. The infectivity of the isolate to wild and laboratory mice was slightly different from that of C. muris. DNA sequences of the 18S ribosomal RNA (rRNA) gene of the isolate were not identical to those of any known Cryptosporidium species. However, phylogenetic analysis indicated that the isolate was a member of the C. muris cluster with minor sequence differences. Therefore, the authors proposed the isolate as a novel genotype of C. muris and named it the C. muris Japanese field mouse genotype (Hikosaka and Nakai, 2005). The third species definitively identified in wild mammals is C. canis, which has been detected in foxes and coyotes by Zhou et al. (2004). In addition to these three established species, 1 genotype of C. hominis (monkey genotype), 2 genotypes of C. canis (fox and coyote genotypes), and nearly 20 host-adapted genotypes of Cryptosporidium (bear, cervine, deer, deer-mouse, ferret, fox, muskrat I and II, marsupial I and II, mouse, mongoose, opossum I and II, rabbit, squirrel A, B, and C, wildebeest, seal I and II, and skunk genotypes) have been identified in wild mammals (Abe and Iseki, 2003; Abe et al., 2004; Xiao et al., 2004; Power et al., 2004; Xiao and Ryan, 2004; Atwill et al., 2004; Alves et al., 2005; Power et al., 2005). In a study undertaken to characterize the Cryptosporidium isolates originating from the naturally infected woodland and field rodents C. glareolus, A. flavicollis, and M. arvalis, Bednarska et al. (2003) found that the measurements of oocyst dimensions and oocyst morphology did not allow distinction between the parasite isolates from the three rodent species. The three groups of isolates produced significantly different pictures of infection in C57BL/6 mice. The successful transmission from wild hosts to laboratory rodents, and the close similarity of the Cryptosporidium oocyst wall protein (COWP) sequence among these three isolates and the “mouse” genotype, and between the “mouse” genotype and C. parvum, indicate that small rodents should be considered an important reservoir of Cryptosporidium genotypes closely related to C. parvum and potentially hazardous for human health. In a study performed at the zoo of Lisbon, Portugal, the Cryptosporidium mouse genotype was detected in an asymptomatic Prairie bison (Bison bison bison) (Alves et al., 2005). The mouse genotype is very common in mice and
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small rodents (Morgan et al., 1999; Bajer et al., 2003) and, phylogenetically, it has the closest relatedness to C. parvum, which infects mainly ruminants (Xiao et al., 2004). The authors postulated that the bison was infected with the mouse genotype. However, the absence of clinical signs, the low level of oocyst shedding, and its detection only once in one animal could not exclude the possibility of the mere mechanical passage of oocysts through the gastrointestinal tract of the animal. In the same study at the Lisbon Zoo (Alves et al., 2005), the authors identified a new Cryptosporidium genotype in the feces of an asymptomatic black wildebeest. Due to its similarity with the W5 genotype from storm water, the authors thought the new genotype could represent a new Cryptosporidium parasite that the black wildebeest acquired from other animals in the zoo, rather than a host-specific parasite from Africa. Cryptosporidium infection is apparently common in wildebeests. An earlier study identified Cryptosporidium in black wildebeests in the Mikumi National Park, Morogoro, Tanzania (Mtambo et al., 1997), but molecular characterization was not done. Cryptosporidium was also previously found in feces of a blue wildebeest (Connochaetes taurinus taurinus) at the Barcelona zoo (Gómez et al., 2000). It was molecularly characterized as C. parvum (Morgan et al., 1999a). Thus, similar to many species or groups of vertebrates (Xiao et al., 2004), wildebeests may be infected with more than one species of Cryptosporidium, C. parvum, and this new Cryptosporidium genotype. Many other wild mammals have been reported to be susceptible to multiple species or genotypes. Deer are susceptible to C. parvum, Cryptosporidium cervine genotype, and the Cryptosporidium deer genotype (Ryan et al., 2003; Da Silva et al., 2003; Xiao et al., 2004). Bactrian camels are susceptible to C. andersoni and C. muris (Xiao et al., 2004). Foxes have been found infected with Cryptosporidium fox genotype, C. canis fox genotype, C. canis dog genotype, and Cryptosporidium muskrat genotype (Xiao et al., 2002; Zhou et al., 2004). Kangaroos have been found infected with three types of Cryptosporidium—the marsupial genotypes I and II, and genotype EGK1 (Power et al., 2004). Opossums have been found infected with Cryptosporidium opossum genotypes I and II (Xiao et al., 2004). Mice have been found infected with C. muris and the Cryptosporidium mouse genotype (Xiao et al., 2004). Muskrats are susceptible to Cryptosporidium muskrat genotypes I and II (Zhou et al., 2004). Squirrels have been found infected with at least three types of Cryptosporidium: genotypes CGS-A, CGS-B, and CGS-C (Atwill et al., 2004).
IV.
Sources of Cryptosporidium Infection and Transmission Dynamics
Molecular tools, besides helping to clarify the taxonomy of Cryptosporidium, are very helpful in assessing the zoonotic potential of various Cryptosporidium species in wild mammals and sources of human and animal infections, and are playing a significant role in the characterization of Cryptosporidium transmission dynamics. The significance of wild animals as reservoirs of Cryptosporidium species or genotypes pathogenic for humans and domestic animals, possibly as a source of contamination of water supplies, is controversial. The occurrence of human-pathogenic Cryptosporidium species in some wild mammals and Cryptosporidium genotypes from wildlife in source water (Xiao et al., 2000; Jellison et al., 2002; Xiao et al., 2006) demonstrates the potential for transmitting the infection from wildlife to farm animals and to water supplies. In addition, some wild animals, particularly rodents and raccoon dogs, can be found in both urban and rural environments, increasing the opportunity for transmission of C. parvum and C. muris infections to humans and domestic animals (Katsumata et al., 2000; Gatei et al., 2003; McGuigan, 2005). It has been speculated that wild animals living within the confines of watercourses used for human consumption, or even aquatic rodents such as muskrats, can play a role in the transmission of zoonotic species and genotypes of Cryptosporidium that can affect humans, livestock, domestic pets, or species kept in captivity, by surface water contamination (Quy et al., 1999; Perz and Blancq, 2001). All these suggestions are largely based on the assumption that these animals are infected with C. parvum, but this supposition is not supported by molecular studies. A few studies have been conducted to determine the importance of domestic animals and terrestrial wildlife in the contamination of watersheds. In Canada and Australia, the authors found that concentrations of Cryptosporidium oocysts in wildlife feces were significantly lower than those in domestic
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livestock or human effluent, indicating that emphasis should be placed essentially on domestic animal management (Heitman et al., 2002; Cox et al., 2005). The strong host adaptation exhibited by the majority of species and genotypes of Cryptosporidium could mean that these parasites do not have high infectivity for humans. In fact, the majority of genotypes found in wild animals are found in a restricted group of hosts. In the United States, five species of mammals that inhabited wetlands adjacent to a fluvial zone (foxes, raccoons, muskrats, otters, and beavers) were examined for cryptosporidial infection (Zhou et al., 2004). Molecular characterization of oocysts found in these hosts led to identification of C. canis, C. canis fox genotype, and three more host-adapted genotypes of Cryptosporidium, which have not been found in humans or domestic animals. The authors concluded that host-adapted species and genotypes of Cryptosporidium excreted by these animals do not pose a threat to public health. Nevertheless, strict host specificity is not certain. It is not uncommon to find one species or genotype of Cryptosporidium in multiple hosts. For example, consider the host ranges of C. parvum, C. meleagridis, C. felis, C. canis, C. suis, C. muris, and the Cryptosporidium cervine genotype. Except for C. parvum and C. meleagridis, which are found frequently in humans and infect a large number of other hosts, the remaining species infect a limited number of hosts and have been associated with a small number of human cases, mostly children and human immunodeficiency virus (HIV) patients. The possibility of transmission of these species to humans through domestic and wild animals cannot be excluded. Although the specific role of wild mammals in the transmission of Cryptosporidium to humans is unknown, it is important for the scientific community and public health authorities to be aware that less prevalent species and genotypes, not yet found in humans, could emerge as human pathogens (Xiao et al., 2002, 2004; Xiao and Ryan, 2004). We are perhaps now experiencing the emergence of the Cryptosporidium cervine genotype as an important human pathogen, which infects various species of wild mammals, is widely seen in surface water, and is being found in increasing numbers of human cases.
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Morgan, U.M, Sturdee, A.P., Singleton, G., Gómez, M.S, Gracenea, M., Torres, J., Hamilton, S.G., Woodside, D.P. and Thompson, R.C.A. 1999. The Cryptosporidium ‘‘mouse’’ genotype is conserved across geographic areas. J. Clin. Microbiol. 37, 1302–1305. Morgan, U.M., Xiao, L., Fayer, R., Lal, A.A. and Thompson, R.C.A. 1999a. Variation in Cryptosporidium: towards a taxonomic revision of the genus. Int. J. Parasitol. 29, 1733–1751. Morgan, U.M., Xiao, L., Monis, P., Sulaiman, I., Pavlasek, I., Blagburn, B., Olson, M., Upton, S.J., Khramtsov, N.V., Lal, A., Elliot, A. and Thompson, R.C. 2000. Molecular and phylogenetic analysis of Cryptosporidium muris from various hosts. Parasitology. 120, 457–464. Mosier, D.A., Cimon, K.Y., Kuhls, T.L., Oberst, R.D. and Simons, K.R. 1997. Experimental cryptosporidiosis in adult and neonatal rabbits. Vet. Parasitol. 69, 163–169. Mtambo, M.M., Nash, A.S., Blewett, D.A, Smith, H.V. and Wright, S. 1991. Cryptosporidium infection in cats: prevalence of infection in domestic and feral cats in the Glasgow area. Vet. Rec. 129, 502–504. Mtambo, M.M., Nash, A.S., Wright, S.E., Smith, H.V., Blewett, D.A. and Jarrett, O. 1995. Prevalence of specific anti-Cryptosporidium IgG, IgM, and IgA antibodies in cat sera using an indirect immunofluorescence antibody test. Vet. Parasitol. 60, 37–43. Mtambo, M.M., Sebatwale, J.B., Kambarage, D.M., Muhairwa, A.P., Maeda, G.E., Kusiluka, L.J.M. and Kazwala, R.R. 1997. Prevalence of Cryptosporidium spp. oocysts in cattle and wildlife in Morogoro region, Tanzania. Prev. Vet. Med. 31, 185–190. Nakai, Y., Hikosaka, K., Sato, M., Sasaki, T., Kaneta, Y. and Okazaki, N. 2004. Detection of Cryptosporidium muris type oocysts from beef cattle in a farm and from domestic and wild animals in and around the farm. J. Vet. Med. Sci. 66, 983–984. Nizeyi, J.B., Cranfield, M.R. and Graczyk, T.K. 2002. Cattle near the Bwindi Impenetrable National Park, Uganda, as a reservoir of Cryptosporidium parvum and Giardia duodenalis for local community and free-ranging gorillas. Parasitol. Res. 88, 380–385. Nizeyi, J.B., Mwebe, R., Nanteza, A., Cranfield, M.R., Kalema, G.R. and Graczyk, T.K. 1999. Cryptosporidium sp. and Giardia sp. infections in mountain gorillas (Gorilla gorilla beringei) of the Bwindi Impenetrable National Park, Uganda. J. Parasitol. 85, 1084–1088. Nizeyi, J.B., Sebunya, D., Da Silva, A.J., Cranfield, M.R., Pieniazek, N.J. and Graczyk, T.K. 2002a. Cryptosporidiosis in people sharing habitats with free-ranging mountain gorillas (Gorilla gorilla beringei), Uganda. Am. J. Trop. Med. Hyg. 66, 442–444. Nutter, F.B., Dubey, J.P., Levine, J.F., Breitschwerdt, E.B, 2004. Seroprevalences of antibodies against Bartonella henselae and Toxoplasma gondii and fecal shedding of Cryptosporidium spp. Giardia spp. and Toxocara cati in feral and pet domestic cats. J. Am. Vet. Med. Assoc. 225, 1394–1398. O’Donoghue, P.J. 1995. Cryptosporidium and cryptosporidiosis in man and animals. Int. J. Parasitol. 25, 139–195. Olson, M.E., O’Handley, R.M., Ralston, B.J., McAllister, T.A. and Thompson, R.C. 2004. Update on Cryptosporidium and Giardia infections in cattle. Trends Parasitol. 20, 185–191. Peeters, J.E., Geeroms, R., Carman, R.J. and Wilkins, T.D. 1986. Significance of Clostridium spiroforme in the enteritis-complex of commercial rabbits. Vet. Microbiol. 12, 25–31. Perz, J.F. and Le Blancq, S.M. 2001.Cryptosporidium parvum infection involving novel genotypes in wildlife from lower New York State, Appl. Environ. Microbiol. 67, 1154–1162. Pospischil, A., Stiglmair-Herb, M.T., Hegel, G.Von. and Wiesner, H. 1987. Abomasal cryptosporidiosis in mountain gazelles. Veterinary Record. 121, 379–380. Power, M.L., Slade, M.B., Sangster, N.C. and Veal, D.A. 2004. Genetic characterisation of Cryptosporidium from a wild population of eastern grey kangaroos Macropus giganteus inhabiting a water catchment. Infect. Genet. Evol. 4, 59–67. Power, M.L., Sangster, N.C., Slade, M.B. and Veal, D.A. 2005. Pattern of Cryptosporidium oocysts shedding by eastern grey kangaroos inhabiting an Australian watershed. Appl. Environ. Microbiol. 71, 6159–6164. Quy, R.J., Cowan, D.P., Haynes, P.J., Sturdee, A.P., Chalmers, R.M., Bodley-Tickell, A.T. and Bull, S.A. 1999. The Norway rat as a reservoir host of Cryptosporidium parvum. J. Wildl. Dis. 35, 660–670. Ramirez, N.E., Ward, L.A. and Sreevatsan, S.A. 2004. Review of the biology and epidemiology of cryptosporidiosis in humans and animals. Microbes Infect. 6, 773–785. Rehg, J.E., Lawton, G.W. and Pakes, S.P. 1979. Cryptosporidium cuniculus in the rabbit (Oryctolagus cuniculus). Lab. Anim. Sci. 29, 656–660.
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Rickard, L.G., Siefker, C., Boyle, C.R. and Gentz, E.J. 1999. The prevalence of Cryptosporidium and Giardia spp. in fecal samples from free-ranging white-tailed deer (Odocoileus virginianus) in the southeastern United States. J. Vet. Diagn. Invest. 11, 65–72. Riggs, M.W. 1990. Cryptosporidiosis in cats, dogs, ferrets, raccoons, opossums, rabbits, and nonhuman primates, in Cryptosporidiosis in Man and Animals, Dubey, J.P., Speer, C.A. and Fayer, R., eds., CRC Press, Boca Raton, Florida. 113–123. Ryan, M.J., Sundberg, J.P., Sauerschell, R.J. and Todd, K.S. Jr. 1986. Cryptosporidium in a wild cottontail rabbit (Sylvilagus floridanus). J. Wildl. Dis. 22, 267. Ryan, U., Xiao, L., Read, C., Zhou, L., Lal, A.A. and Pavlasek, I. 2003. Identification of novel Cryptosporidium genotypes from the Czech Republic. Appl. Environ. Microbiol. 69, 4302–4307. Ryan, U.M., Monis, P., Enemark, H.L., Sulaiman, I., Samarasinghe, B., Read, C., Buddle, R., Robertson, I., Zhou, L., Thompson, R.C. and Xiao, L. 2004. Cryptosporidium suis n. sp (Apicomplexa: Cryptosporidiidae) in pigs (Sus scrofa). J. Parasitol. 90, 769–773. Siefker, C., Rickard, L.G., Pharr, G.T., Simmons, J.S., O’Hara, T.M. 2002. Molecular characterization of Cryptosporidium sp. isolated from northern Alaskan caribou (Rangifer tarandus). J. Parasitol. 88, 213–216. ´ Sinski, E., Bednarska, M. and Bajer A. 1998. The role of wild rodents in ecology of cryptosporidiosis in Poland. Folia Parasitol. 45, 173 and 174. ´ Sinski, E., Hlebowicz, E. and Bednarska, M. 1993. Occurrence of Cryptosporidium parvum infection in wild small mammals in District of Mazury Lake (Poland). Acta Parasitol. 38, 59–61. Snyder, D.E. 1988. Indirect immunofluorescent detection of oocysts of Cryptosporidium parvum in the feces of naturally infected raccoons (Procyon lotor). J. Parasitol. 74, 1050–1052. Sturdee, A.P., Bodley-Tickell, A.T., Archer, A. and Chalmers, R.M. 2003. Long-term study of Cryptosporidium prevalence on a lowland farm in the United Kingdom. Vet. Parasitol. 116, 97–113. Sturdee, A.P., Chalmers, R.M. and Bull, S.A. 1999. Detection of Cryptosporidium oocysts in wild mammals of mainland Britain. Vet. Parasitol. 80, 273–280. Sundberg, J.P., Hill, D. and Ryan, M.J. 1982. Cryptosporidiosis in a gray squirrel. J. Am. Vet. Med. Assoc. 181, 1420. Tarazona, R., Blewett, D.A. and Carmona M.D. 1998. Cryptosporidium parvum infection in experimentally infected mice: infection dynamics and effect of immunosuppression. Folia Parasitol. (Praha). 45, 101–107. Torres, J., Gracenea, M., Gomez, M.S., Arrizabalaga, A. and Gonzalez-Moreno, O. 2000. The occurrence of Cryptosporidium parvum and C. muris in wild rodents and insectivores in Spain. Vet. Parasitol. 92, 253–260. Tyzzer, E.E. 1907. A sporozoan found in the peptic glands of the common mouse. Proc. Soc. Exp. Biol. Med. 5, 12 and 13. Tyzzer, E.E. 1910. An extracellular coccidium Cryptosporidium muris (gen. et sp. nov.) of the gastric glands of the common mouse. J. Med. Res. 23, 487–509. Tyzzer, E.E. 1912. Cryptosporidium parvum (sp. nov.), a coccidium found in the small intestine of the common mouse. Arch. Protistenkd. 26, 394–412. Tzipori, S. and Campbell, I. 1981. Prevalence of Cryptosporidium antibodies in 10 animal species. J. Clin. Microbiol. 14, 455 and 456. Upton, S.J. and Current, W.L. 1985. The species of Cryptosporidium (Apicomplexa: Cryptosporidiidae) infecting mammals. J. Parasitol. 71, 625–629. Van Winkle, T.J. 1985. Cryptosporidiosis in young artiodactyls. J. Am. Vet. Med. Assoc. 187, 1170–1172. Warren, K.S., Swan, R.A., Morgan-Ryan, U.M., Friend, J.A. and Elliot, A. 2003. Cryptosporidium muris infection in bilbies (Macrotis lagotis). Aust. Vet. J. 81, 739–741. Webster, J.P. and MacDonald, D.W. 1995. Cryptosporidiosis reservoir in wild brown rats (Rattus norvegicus) in the UK. Epidemiol. Infect. 115, 207–209. Xiao, L. and Ryan, U.M. 2004. Cryptosporidiosis: an update in molecular epidemiology. Curr. Opin. Infect. Dis. 17, 483–490. Xiao, L., Alderisio, K., Limor, J., Royer, M. and Lal, A.A. 2000. Identification of species and sources of Cryptosporidium oocysts in storm waters with a small subunit rRNA-based diagnostic and genotyping tool. Appl. Environ. Microbiol. 66, 5492–5498.
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Xiao, L., Alderisio, K. and Jiang, J. 2006. Detection of Cryptosporidium oocysts in water: effect of the number of samples and analytic replicates on test results. Appl. Environ. Microbiol. 72, 5942–5947. Xiao, L., Escalante, L., Yang, C., Sulaiman, I., Escalante, A.A., Montali, R.J., Fayer, R. and Lal, A.A. 1999. Phylogenetic analysis of Cryptosporidium parasites based on the small-subunit rRNA gene locus. Appl. Environ. Microbiol. 65, 1578–1583. Xiao, L., Fayer, R., Ryan, U. and Upton, S.J. 2004. Cryptosporidium taxonomy: recent advances and implications for public health, Clin. Microbiol. Rev. 17, 72–97. Xiao, L., Sulaiman, I.M., Ryan, U.M., Zhou, L., Atwill, E.R., Tischler, M.L., Zhang, X., Fayer, R. and Lal, A.A. 2002. Host adaptation and host-parasite coevolution in Cryptosporidium: implications for taxonomy and public health. Int. J. Parasitol. 32, 1773–1785. Yamura, H., Shirasaka, R., Asahi, H., Koyama, T., Motoki, M. and Ito, H., 1990. Prevalence of Cryptosporidium infection among house rats, Rattus rattus and R. norvegicus, in Tokyo, Japan and experimental cryptosporidiosis in roof rats. Jpn. J. Parasitol. 39, 439–444. Zhou, L., Fayer, R., Trout, J.M., Ryan, U.M., Schaefer, F.W. 3rd and Xiao, L. 2004. Genotypes of Cryptosporidium species infecting fur-bearing mammals differ from those of species infecting humans. Appl. Environ. Microbiol. 70, 7574–7577.
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17 Companion Animals
Mónica Santín and James M. Trout
CONTENTS I. Introduction .................................................................................................................................. 437 II. Cryptosporidiosis in Cats ............................................................................................................. 437 III. Cryptosporidiosis in Dogs............................................................................................................ 440 IV. Cryptosporidiosis in Horses ......................................................................................................... 442 References........................................................................................................................... 444
I.
Introduction
Pet ownership, particularly of dogs and cats, is common throughout the world. Gastrointestinal disorders in companion animals, especially diarrhea, are a common reason for pet owners to seek for veterinary advice. There is a modest prevalence of Cryptosporidium sp. infections in companion animals. Although clinical disease is observed infrequently (e.g., Mtambo et al., 1991; Nash et al., 1993; Xiao and Herd, 1994a; Abe et al., 2002a; Ederli et al., 2005), severe clinical illness has been reported in dogs, cats, and horses, occasionally associated with concurrent infections such as canine distemper virus, isosporiasis, or feline leukemia virus (e.g., Snyder et al., 1978; Monticello et al., 1987; Goodwin and Barsanti, 1990; Miller et al., 2003; Aydin et al., 2004). Although pets offer significant benefits to society, there are documented health hazards associated with owning a pet, including the potential risk to humans of enteric parasites harbored by companion animals. Cryptosporidiosis detected in dogs, cats, and horses could represent an important reservoir of infection for humans. Because of the possible zoonotic implication, veterinarians need to recognize this potential in pets and provide accurate advice to their clients. This is of particular importance to highrisk populations, such as children, the elderly, and the immunocompromised.
II.
Cryptosporidiosis in Cats
Cryptosporidium sp. was first reported in cats in Japan (Iseki, 1979). Sporulated oocysts (5.0 × 4.5 µm) were found in the feces of naturally infected cats. Following observations of the endogenous stages and transmission and cross-transmission studies, the organism was named C. felis (Iseki, 1979). Subsequently, asymptomatic and symptomatic Cryptosporidium infections have been reported in cats worldwide (Bennett et al., 1985; Arai et al., 1990; Mtambo et al., 1991; Lappin et al., 1997a; Morgan et al., 1998; Hill et al., 2000; Ryan et al., 2003; Cirak and Bauer, 2004; Fayer et al., 2006; Santín et al., 2006). Seroprevalence studies conducted in Scotland (Tzipori and Campbell, 1981; Mtambo et al., 1995), Germany (Cirak and Bauer, 2004), the United States (Lappin et al., 1997a; McReynolds et al., 1999; Nutter et al., 2004), and the Czech Republic (Svobodova et al., 1994) reported that 8.3 to 74% of cats
437
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Cryptosporidium and Cryptosporidiosis, Second Edition TABLE 17.1 Prevalence of Cryptosporidium spp. Reported in Cats by Microscopy Country
Number of Animals
Prevalence (% positive)
Reference
Australia Australia Czech Republic Germany Germany Germany Japan Japan Scotland Scotland United States United States United States United States
418 162 135 200 100 441 608 507 235 57 205 170 176 263
0 1.2 5.9 0 1 0 3.8 3.9 8.1 12.3 5 2.4 7 3.8
McGlade et al., 2003 Sargent et al., 1998 Svobodova et al., 1994 Augustin-Bichl et al., 1984 Cirak and Bauer, 2004 Epe et al., 2004 Arai et al., 1990 Uga et al., 1989 Mtambo et al., 1991 Nash et al., 1993 Hill et al., 2000 Lappin et al., 1997a Nutter et al., 2004 Spain et al., 2001
surveyed had detectable antibodies to Cryptosporidium spp. However, detection of Cryptosporidiumspecific IgG in cat serum only suggests prior infection and does not correlate with active excretion of oocysts (Mtambo et al., 1995; Lappin et al., 1997a; Cirak and Bauer, 2004). Conventional coproscopical methods used to detect Cryptosporidium oocysts in studies in Australia, the Czech Republic, Germany, Japan, Scotland, and the United States found that 0 to 12.3% of cats were excreting oocysts (Table 17.1). However, conventional microscopy, commonly used to identify parasites in fecal samples, is less sensitive than molecular detection methods (Scorza et al., 2003). For example, 418 fecal specimens from Australian cats were found to be negative for Cryptosporidium oocysts by microscopy. However, when 40 randomly selected samples were retested by PCR, 10% were positive (McGlade et al., 2003). PCR was also more sensitive than immunofluorescence microscopy (IFA) in detecting infections during an outbreak in a cat colony (Fayer et al., 2006). In the first 10 days of the outbreak, specimens were examined by both methods, and Cryptosporidium sp. was detected in 92 and 51% of the samples by PCR and IFA, respectively. In addition to greater sensitivity, molecular methods also allow the accurate identification of the species or genotypes of Cryptosporidium that are present in fecal specimens. There is little information available on the prevalence of Cryptosporidium species/genotypes in cats, and genotyping has been conducted on only 35 cat isolates (Table 17.2). Three species of Cryptosporidium have been reported in cats: C. parvum, C. felis, and C. muris (Morgan et al., 1998; Sargent et al., 1998; Alves et al., 2001; Scorza et ˇ et al., 2004; Fayer et al., 2006; Pavlasek and Ryan, 2007; Santín et al., 2006). al., 2003; Hajdusek However, only C. felis and C. muris have been confirmed by molecular analysis in naturally infected cats. Although C. parvum has been reported in cats, identification was based on conventional microscopy and oocyst size, not molecular characterization (Lappin et al., 1997b; Hill et al., 2000). Nevertheless, experimental infection with C. parvum was demonstrated in eight adult domestic cats (Scorza et al., 2003). Sargent et al. (1998) originally suggested the existence of a cat-adapted strain or species (Cryptosporidium feline genotype) based on the sequence analysis of SSU-rRNA gene and oocyst size from naturally infected cats in Australia. Morphological studies revealed oocysts with an average size of 4.6 × 4.0 µm, smaller in size than typical oocysts seen in human stools (5.0 × 4.5 µm), and phylogenetic analysis placed the cat isolates into a distinct group, separate from other Cryptosporidium species (Sargent et al., 1998). Later, Morgan et al. (1998) isolated oocysts of the same size and gene sequences from four cats and referred to them as C. felis, the name already used by Iseki (1979). The validity of C. felis (Iseki, 1979) was in doubt for some years, but recent molecular characterization of different loci supports the concept of C. felis as a valid species (Xiao et al., 2004). More recently, C. felis has been reported
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TABLE 17.2 Cryptosporidium Species/Genotypes Reported in Cats with Molecular Data Cryptosporidium Species/Genotype (Number of Isolates) C. C. C. C.
felis felis felis felis
(2) (4) (1) (2)
C. C. C. C. C.
felis (1) felis (18) felis (5) muris (1) muris (1)
Country
Oocyst Data (µm)
Gene(s)
Australia Australia Portugal Czech Republic
4.6 × 4.0 Same as Sargent et al., 1998 N/A N/A
Czech Republic United States Colombia Colombia Czech Republic
4.6 ± 0.3 × 4.2 ± 0.2 4.6 × 4.2 N/A N/A 8.0 × 4.8
SSU-rRNA SSU-rRNA SSU-rRNA SSU-rRNA HSP-70 SSU-rRNA SSU-rRNA SSU-rRNA SSU-rRNA SSU-rRNA
Reference Sargent et al., 1998 Morgan et al., 1998 Alves et al., 2001 Ryan et al., 2003 ˇ et al., 2004 Hajdusek Fayer et al., 2006 Santín et al., 2006 Santín et al., 2006 Pavlasek and Ryan, 2007
in naturally infected cats in Portugal, the United States, Colombia, and the Czech Republic (Alves et ˇ et al., 2004; Fayer et al., 2006; Santín et al., 2006). In crossal., 2001; Ryan et al., 2003; Hajdusek transmission studies, oocysts from naturally infected cats failed to infect mice, rats, guinea pigs, and dogs (Iseki, 1979; Asahi et al., 1991). However, two lambs were infected with oocysts isolated from a farm cat naturally infected with an unidentified species of Cryptosporidium (Mtambo et al., 1996). Cryptosporidium felis has also been identified in a cow based on oocyst measurements and gene sequence data (Bornay-Llinares et al., 1999). Of 46 cats from Bogota (Colombia), 5 were infected with C. felis and 1 with C. muris (Santín et al., 2006). This was the first report of C. muris in cats; however, it was not possible to determine if the cat simply harbored oocysts from an infected rodent it had eaten or if it had an active infection. An additional natural C. muris infection has been reported in a cat with gastroenteritis from the Czech Republic (Pavlasek and Ryan, 2007). In this cat, six out of the seven fecal specimens analyzed during a 1-month period were positive for Cryptosporidium, suggesting that this was an active natural infection. Moreover, an experimental infection with C. muris in three cats has been reported, with large numbers of developing parasites observed in mucosal scrapings from the stomach (Iseki et al., 1989). Most studies of experimentally and naturally infected cats reported shedding of Cryptosporidium oocysts without the presence of clinical signs (Augustin-Bichl et al., 1984; Arai et al., 1990; Asahi et al., 1991; Mtambo et al., 1991; Nash et al., 1993; Fayer et al., 2006). However, oocysts have been observed in the feces of cats with persistent diarrhea (Barr et al., 1994; Bennett et al., 1985; Goodwin and Barsanti, 1990; Lent et al., 1993; Monticello et al., 1987; Morgan et al., 1998). The incidence and clinical importance of cryptosporidiosis in cats are unknown, but cryptosporidiosis might be included in the differential diagnosis of chronic feline diarrhea. There are no known effective methods for chemical prophylaxis or treatment of feline cryptosporidiosis. In only one case was paramomycin effective in clearing Cryptosporidium oocysts from feces from a cat with persistent diarrhea (Barr et al., 1994). Cryptosporidium felis has been implicated as sources of cryptosporidial infections in both immunocompromised and immunocompetent humans in several countries (Pedraza-Diaz et al., 2001; Xiao et al., 2001; Cacciò et al., 2002; Pieniazek et al., 1999; Gatei et al., 2002, 2006; Guyot et al., 2001; McLauchlin et al., 2000; Morgan et al. 2000a; Cama et al., 2003; Matos et al., 2004; Leoni et al., 2006). Morgan et al. (2000a) noted that three of six patients infected with C. felis had pet cats, and in England a cat and his owner were concurrently infected with Cryptosporidium sp. (Bennett et al., 1985). However, it was not possible to determine whether one infected the other or if both were infected from a common source. When attempting to determine the source of cryptosporidiosis in humans, contact with companion animals should be considered.
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III. Cryptosporidiosis in Dogs The first evidence of cryptosporidiosis in dogs was reported by Tzipori and Campbell (1981), who detected Cryptosporidium antibodies in 16 of 20 canine serum samples. The first clinical case was reported 2 years later when life-cycle stages characteristic of Cryptosporidium sp. were identified in a 1-week-old puppy with acute diarrhea (Wilson et al., 1983). Since then, cryptosporidiosis in dogs has been reported worldwide, involving both asymptomatic and diarrheic dogs (e.g., Greene et al., 1990; Johnston and Gasser, 1993; Causapé et al., 1996; Abe et al., 2002a; Hackett and Lappin, 2003; Cirak and Bauer, 2004; Ederli et al., 2005; Giangaspero et al., 2006). Large-scale surveys of Cryptosporidium infection in dogs have been performed in several countries using different diagnostic methods. Coprological studies by microscopy found infection rates ranging from 0.23 to 40% in Argentina, Australia, Brazil, the Czech Republic, Egypt, Germany, Hungary, India, Japan, Korea, Spain, and the United States (see Table 17.3). In contrast, no Cryptosporidium oocysts were observed in fecal specimens from 421 dogs surveyed in Australia (Bugg et al., 1999), 1281 and 200 dogs studied in two different surveys in Germany (Augustin-Bichl et al., 1984; Epe et al., 2004), 101 dog admitted to kennels in Scotland (Simpson et al., 1988), or 57 dogs in an international dog show in Finland (Pohjola, 1984). A commercial Cryptosporidium coproantigen ELISA test (ProSpecT Cryptosporidium Microplate Assay, Alexon, Inc., Sunnyvale, CA) was more sensitive than conventional microscopical methods for 270 dogs examined using both methods (23 and 0.4%, respectively) (Cirak and Bauer, 2004). However, the authors were concerned about false-positives and recommended confirming a positive ELISA by fecal examination using another detection method. TABLE 17.3 Prevalence of Cryptosporidium spp. Reported in Dogs by Microscopy Country
No. of Animals
Prevalence (% positive)
Reference
Argentina Argentina Australia Australia Australia Brazil Brazil Brazil Brazil Czech Republic Egypt Egypt Finland Germany Germany Hungary India Japan Japan Korea Scotland Spain United States United States United States United States
2193 113 421 493 55 100 166 450 433 458 685 25 57 270 1281 36 9 140 213 257 101 81 200 130 100 49
0.23 12.38 0 11 1.8 40 2.41 8.8 1.4 4.6 3.8 12 0 0.4 0 8.3 8.53 6.4 1.4 9.7 0 7.4 2 3.8 17 10.2
Fontanarrosa et al., 2006 Ponce de León et al., 1994 Bugg et al., 1999 Johnston and Gasser, 1993 Milstein and Goldsmid, 1995 Ederli et al., 2005 Huber et al., 2005 Lallo and Fernandes Bondan, 2006 Mundim et al., 2006 Svobodova et al., 1994 Abou-Eisha and Abdel-Aal, 1995 El-Hohary and Abdel-Latif, 1998 Pohjola, 1984 Cirak and Bauer, 2004 Epe et al., 2004 Nagy, 1995 Kumar et al., 2004 Abe et al., 2002a Uga et al., 1989 Kim et al., 1998 Simpson et al., 1988 Causapé et al., 1996 El-Ahraf et al., 1991 Hackett and Lappin, 2003 Juett et al., 1996 Jafri et al., 1993
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Seroepidemiological surveys found Cryptosporidium infection to be relatively common, with antiCryptosporidium antibodies detected in 266 of 458 dogs (58%) examined in the Czech Republic (Svobodova et al., 1994) and in 16 of 20 dogs (80%) examined in the United States (Tzipori and Campbell, 1981). Comparisons of living conditions and infection status of dogs have shown mixed results: Cryptosporidium prevalence was similar among stray and domestic dogs in Spain (6.8% versus 8.1%) (Causapé et al., 1996), but a higher prevalence was observed in kennel dogs than in privately owned dogs in Italy (5% versus 1.7%) (Giangaspero et al., 2006). The relationship between age of the dogs and infection with Cryptosporidium spp. is not clear. One study reported a higher prevalence in puppies (<1 year) than in adults from the United States (31 and 5%, respectively) (Juett et al., 1996); conversely, in Brazil, a lower prevalence of infection in puppies (5.5%) compared to adults was reported (10.1%) (Lallo and Fernandes Bondan, 2006). Other studies have reported no age difference in the percentage of Cryptosporidium-infected dogs (Huber et al., 2005; Giangaspero et al., 2006). Prevalence of infection with Cryptosporidium spp. was not found to be associated with the sex of the dogs (Uga et al., 1989; El-Ahraf et al., 1991; Juett et al., 1996; Huber et al., 2005; Lallo and Fernandes Bondan, 2006). Most dogs infected with Cryptosporidium spp. have been asymptomatic (Uga et al., 1989; Abe et al., 2002a; Ederli et al., 2005). Clinical signs typically have been reported in puppies with concurrent diseases, such as canine distemper, parvovirus enteritis, or a known immunodeficiency (Wilson et al., 1983; Turnwald et al., 1988; Fukushima and Helman, 1984; Greene et al., 1990; Willard and Bouley, 1999; Miller et al., 2003; Aydin et al., 2004). The presence of concomitant infection makes it difficult to determine if Cryptosporidium spp. are the primary cause of the observed clinical signs. Cryptosporidiosis might be relatively common in dogs, because it can be a difficult disease for clinicians to diagnose due to the small size of the oocysts, the absence of clinical signs, and the intermittent nature of oocyst excretion. Thus, multiple stool samples should be tested before a negative result is reported. Most studies have relied on morphometric identification of the parasite and have not attempted to identify species by molecular methods. Without genetic analysis, it is impossible to accurately determine the species or genotype in a specific isolate of Cryptosporidium. Therefore, there is limited information available about which Cryptosporidium species and genotypes infect dogs. Until now, only 36 isolates obtained from dogs in Australia, the Czech Republic, Italy, Japan, and the United States have been genotyped. Of these samples, 26 were found to be C. canis, 9 were C. parvum, and 1 was C. meleagridis (Table 17.4). Although these three species are known to be pathogenic to humans, C. canis and C. meleagridis have narrow host ranges, primarily infecting dogs and birds, respectively (Xiao et al., 2004). Given the potential public health risk associated with cryptosporidiosis and, thus, the need to determine the species and genotypes of Cryptosporidium within a given sample, more specific and sensitive diagnostic methods should be selected over traditional methods such as microscopy. The dog genotype (Xiao et al., 1999; Morgan et al., 2000b) was designated as a new species, C. canis, in 2001 based of the results of cross-transmission experiments and genomic analysis (Fayer et al., 2001). Experimental inoculation with C. canis oocysts induced infections in three calves (Fayer et al., 2001), but not in BALB/c neonatal mice, SCID mice, or immunosuppressed C57BL6 juvenile mice (Fayer et al., 2001; Satoh et al., 2006). Cryptosporidium canis oocysts are morphologically similar to those of C. parvum, which is why most cases of cryptosporidiosis reported in dogs before the application of molecular techniques were considered to be caused by C. parvum. Thirty cases of infection with C. canis have been reported in asymptomatic immunocompetent and immunocompromised humans in England, Kenya, Peru, Thailand, and the United States (Pieniazek et al., 1999; McLauchlin et al., 2000; Pedraza-Diaz et al., 2001; Xiao et al., 2001; Gatei et al., 2002, 2006; Cama et al., 2003; Leoni et al., 2006). Dogs in the household have been thought to be a risk factor for Cryptosporidium-associated diarrhea in young children (Molbak et al., 1994). Although there are no reports of dogs being the definitive source of oocysts in human infections, this possibility has been inferred from the development of a cryptosporidial infection in a veterinary student who treated a dog with chronic cryptosporidiosis (unknown Cryptosporidium species) (Greene et al., 1990). Dogs, particularly those in homes of immunocompromised humans, should be evaluated for enteric zoonotic agents. Veterinarians need to be aware of the zoonotic potential of Cryptosporidium spp. when advising potentially immunocompromised clients.
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Cryptosporidium and Cryptosporidiosis, Second Edition TABLE 17.4 Cryptosporidium Species/Genotypes Reported in Dogs with Molecular Data Cryptosporidium Species/Genotype (Number of Isolates)
Oocyst Data (µm)
Country
C. canis (1) C. canis (8)
United States Australia
N/A 4.9 × 4.4
C. canis (1)
United States
4.95 × 4.71
C. parvum (1)
United States
5.19 × 4.9
C. C. C. C. C. C. C. C.
Japan Japan USA Czech Republic Czech Republic Italy Italy Japan
N/A N/A N/A 4.7 ± 0.0 × 4.3 ± 0.3 4.7 ± 0.0 × 4.3 ± 0.3 N/A N/A 4.4 ± 0.5 × 3.2 ± 0.4
canis (13) canis (1) canis (1) parvum (1) meleagridis (1) parvum (7) canis (1) canis (1)
Gene(s) SSU-rRNA SSU-rRNA HSP-70 SSU-rRNA HSP-70 SSU-rRNA HSP-70 SSU-rRNA SSU-rRNA Actin SSU-rRNA SSU-rRNA COWP COWP SSU-rRNA
Reference Xiao et al., 1999 Morgan et al., 2000b Fayer et al., 2001 Fayer et al., 2001 Abe et al., 2002a Abe et al., 2002b Miller et al., 2003 ˇ et al., 2004 Hajdusek ˇ et al., 2004 Hajdusek Giangaspero et al., 2006 Giangaspero et al., 2006 Satoh et al., 2006
Note: N/A = not available.
IV.
Cryptosporidiosis in Horses
Data on equine cryptosporidiosis are scarce. Cryptosporidiosis in horses was initially described in five immunodeficient Arabian foals (Snyder et al., 1978). This early finding suggested that an immunodeficient state was a prerequisite to establishment of the infection in horses. Subsequently, however, cryptosporidiosis has been reported in immunocompetent horses worldwide (Gajadhar et al. 1985; Browning et al., 1991; Xiao and Herd, 1994a; Cole et al. 1998; Coleman et al., 1989; Olson et al., 1997; Majewska et al., 1999, 2004; Mahdi and Ali, 2002; Grinberg et al., 2003; Ryan et al., 2003; Sturdee et al., 2003; Epe et al., 2004). The prevalence of Cryptosporidium spp. in fecal specimens from horses has shown wide variation (Table 17.5). Great differences were observed within and between countries. Reported infection rates determined by microscopy varied from 1 to 47% (Table 17.5). In a 5-year survey (1992–1996) of horses in the United Kingdom, the prevalence of Cryptosporidium spp. varied from 1.9% in 1994 to 24.7% in 1992 (Sturdee et al., 2003). The absence of equine cryptosporidiosis has also been reported in several studies (Table 17.5). Cryptosporidium oocysts were not present in any of the fecal specimens examined from 223 horses and 66 mules used as pack stock at commercial or governmental operations, 91 horses used for backcountry recreation, or 311 horses and mules used commercially in California (Johnson et al., 1997; Fio and Atwill, 1998; Atwill et al., 2000). Cryptosporidiosis was not found in 14 quarterhorse foals with diarrhea in Ohio (Reinemeyer et al., 1984) nor in 8 horses in Egypt (Abou-Eisha, 1994). The presence or absence of cryptosporidial infection is diagnosed primarily by the examination of a single fecal sample. This is likely results in underdiagnosis of infections because oocyst excretion in feces is often intermittent (Xiao and Herd, 1994a). Multiple samplings are necessary to accurately determine the presence of Cryptosporidium oocysts in feces with all diagnostic methods. In studies including multiple specimens the prevalence was higher than that reported when only a single specimen was examined for the presence of oocysts. In a study conducted in Ohio, the prevalence in a population of foals based on a single fecal sample was approximately 18%; in the same population the cumulative prevalence was 71% (Xiao and Herd, 1994a). In another longitudinal survey, 546 fecal specimens were collected daily from 22 mixed-breed pony foals; oocysts were found in feces from all foals (Coleman et al., 1989). In the same study, in which fecal specimens were collected biweekly from 14 quarterhorse
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TABLE 17.5 Prevalence of Cryptosporidium spp. Reported in Horses by Microscopy Country
No. of Animals
Prevalence (% positive)
Reference
Britain and Ireland Canada Egypt Germany Germany Iraq Poland Poland United Kingdom United Kingdom 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
403 35 8 37 4399 25 106 318 644 416 223 436 22 14 17 24 10 9 18 31 1331 91 28 222
27 17 0 3 3.5 12 9.4 3.5 47 8.9 0 3 100a 21.4b 11.7c 12.5d 0e 0f 0g 0h 1 0 0 7.2
Browning et al., 1991 Olson et al., 1997 Abou-Eisha, 1994 Beelitz et al., 1996 Epe et al., 2004 Mahdi and Ali, 2002 Majewska et al., 1999 Majewska et al., 2004 Netherwood et al., 1996 Sturdee et al., 2003 Atwill et al., 2000 Cole et al., 1998 Coleman et al., 1989 Coleman et al., 1989 Coleman et al., 1989 Coleman et al., 1989 Coleman et al., 1989 Coleman et al., 1989 Coleman et al., 1989 Coleman et al., 1989 Fio and Atwill, 1998 Johnson et al., 1997 Reinemeyer et al., 1984 Xiao and Herd, 1994a
a
b c
d
e
f
g
h
Cumulative prevalence: 546 fecal samples were collected daily from 22 foals. Cumulative prevalence: 110 fecal samples were collected biweekly 14 foals. Cumulative prevalence: 52 fecal samples were collected sporadically from 17 foals. Cumulative prevalence: 99 fecal samples were collected bi-weekly from 24 foals. Cumulative prevalence: 168 fecal samples were collected daily from 10 foals. Cumulative prevalence: 17 fecal samples were collected sporadically from 9 foals. Cumulative prevalence: 199 fecal samples were collected daily from 18 foals. Cumulative prevalence: 84 fecal samples were collected sporadically from 31 pony foals.
foals (110 specimens) and 24 mixed-breed pony foals (99 specimens), the prevalence of oocysts was 21.4 and 12.5%, respectively (Coleman et al., 1989). Of the diverse methods available to detect Cryptosporidium oocysts in horse feces, the most widely used method has been acid-fast staining of fecal smears. However, acid-fast staining is known to have a low sensitivity and specificity (Cohen and Snowden, 1996). A commercial immunofluorescence stain (MerIFluor, Meridian Diagnostics, Inc., Cincinnati, OH) is at least 10 times more sensitive at detecting Cryptosporidium oocyst than acid-fast staining (Xiao and Herd, 1994b). Likewise, an enzyme immunoassay to detect coproantigens of Cryptosporidium spp. in feces (ProSpecT Cryptosporidium Microplate Assay, Alexon, Inc.) also showed a higher infection rate in 43 horse samples when compared with microscopy (acid-fast staining) (11.6% by microscopy versus 27.9% by immunoassay) (Majewska et al., 1999).
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Foals are thought to be more susceptible to cryptosporidiosis than older animals. Oocysts excretion peaked in foals at 5 to 8 weeks of age in a longitudinal study that included samples from 35 foals from birth until 35 weeks of age (Xiao and Herd, 1994a). The prevalence of Cryptosporidium infection in foals on two farms in Ohio and one farm in Kentucky was 15%, 18%, and 31%, respectively, whereas on the same farms only 1 of 39 weanlings was positive for Cryptosporidium spp., and no oocysts were found in any of the yearlings and mares (Xiao and Herd, 1994a). Conversely, Olson et al. (1997) found a higher prevalence in horses older than 6 months of age than in foals less than 6 months of age (21% versus 10%). The major clinical sign of cryptosporidiosis in both immunodeficient and immunocompetent foals is diarrhea (Coleman et al., 1989; Grinberg et al., 2003), but most Cryptosporidium infections in adult horses are asymptomatic (Majewska et al. 1999, 2004; Sturdee et al., 2003). However, cryptosporidial infection can cause diarrhea in healthy adult horses (McKenzie and Diffay, 2000), and cryptosporidiosis in foals can be asymptomatic (Xiao and Herd, 1994a). Clinical signs in foals apparently vary with age and immune status. Cryptosporidial infection was established by an experimental inoculation of C. parvum oocysts into five young foals with severe combined immunodeficiency (Bjorneby et al., 1991). All foals developed severe persistent watery diarrhea associated with oocyst shedding within 2 to 5 days postinoculation. Lesions in the jejunum and ileum consisted of mild-to-moderate villous atrophy with numerous Cryptosporidium organisms present in the superficial intestinal mucosa. Immunocompetent horses are also susceptible to illness. An outbreak of diarrhea accompanied by shedding of Cryptosporidium-like oocysts in apparently immunocompetent thoroughbred foals was reported in New Zealand (Grinberg et al., 2003). The outbreak affected nine foals that suffered from severe-to-mild disease accompanied by dehydration and weakness. During the outbreak, two foals died from the disease and one was euthanized because of severe morbidity. To date, no specific therapy has proven to be effective in treating Cryptosporidium. Cryptosporidium parvum was considered the etiologic agent reported in most equine infections. However, most reports were based on the size of the oocysts being consistent with that of C. parvum but without molecular confirmation (e.g., Coleman et al. 1989; Beelitz et al., 1996; Majeswska et al., 1999). Only four published reports include gene sequence information to determine the Cryptosporidium species/genotype present in specimens from horses. In the first three reports C. parvum was identified ˇ et al., 2004) and in the fourth, C. parvum and a (Fio and Atwill, 1998; Grinberg et al., 2003; Hajdusek novel Cryptosporidium horse genotype were reported (Ryan et al. 2003). This horse genotype was identified in a 161-day-old Prezwalski’s wild horse in the Czech Republic. Cryptosporidium parvum oocysts shed by infected horses are potentially infectious for humans. There are two reports that relate horse and human cases of cryptosporidiosis to horses. In one case, a veterinary student acquired cryptosporidiosis following exposure to infected foals (Cohen and Snowden, 1997), and in the other, Cryptosporidium oocysts were detected in a fecal sample from a rider who had sporadic contact with infected horses (Majewska et al., 1999). More extensive evaluation, using molecular techniques is necessary to determine the species/genotypes infecting horses and those implicated in severe cryptosporidiosis leading to morbidity and mortality.
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Juett, B.W., Otero, R.B. and Bischoff, W.H. 1996. Cryptosporidium parvum in the domestic dog population of central Kentucky. Trans. Ky. Acad. Sci. 57: 18–21. Kim, J.T., Wee, S.H. and Lee, C.G. 1998. Detection of Cryptosporidium oocysts in canine fecal samples by immunofluorescence assay. Kor. J. Parasitol. 36: 147–149. Kumar, D., Sreekrishnan, R. and Das, S. S. 2004. Cryptosporidiosis in man and animals in Pondicherry. Ind. J. Anim. Sci. 74: 261–263. Lallo, M.A. and Fernandes Bondan, E.F. 2006. Prevalence of Cryptosporidium sp. in institutionalized dogs in the city of Sãn Paulo, Brazil. Rev. Saúde Pública 40: 120–125. Lappin, M.R., Ungar, B., Brown-Hahn, B., Cooper, C.M., Spilker, M., Thrall, M., Hill, S.L., Cheney, J. and Taton-Allen, G. 1997a. Enzyme-linked immunosorbent assay for the detection of Cryptosporidium parvum IgG in the serum of cats. J. Parasitol. 83: 957–960. Lappin, M. Dowers, R. K., Edsell, D., Taton-Allen, G. and Cheney, J. 1997b. Cryptosporidiosis and inflammatory bowel disease in a cat. Feline Pract. 25: 10–13. Lent, S.F., Burkhardt, J.E. and Bolka, D. 1993. Coincident enteric cryptosporidiosis and lymphosarcoma in a cat with diarrhea. J. Am. Anim. Hosp. Assoc. 29: 492–496. Leoni, F., Amar, C., Nichols G., Pedraza-Díaz, S. and McLauchlin, J. 2006. Genetic analysis of Cryptosporidium from 2414 humans with diarrhea in England between 1985 and 2000. J. Med. Microbiol. 55: 703–707. Mahdi, N.K. and Ali, N.H. 2002. Cryptosporidiosis among animal handlers and their livestock in Basrah, Iraq. East Afr. Med. J. 79: 550–553. Majewska, A.C., Werner, A. Sulima, P. and Luty, T. 1999. Survey on equine cryptosporidiosis in Poland and the possibility of zoonotic transmission. Ann. Agric. Environ. Med. 6: 161–165. Majewska, A.C., Solarczyk, P., Tamang, L. and Graczyk, T.K. 2004. Equine Cryptosporidium parvum infections in western Poland. Parasitol. Res. 93: 274–278. Matos, O., Alves, M., Xiao, L., Cama, V. and Antunes, F. 2004. Cryptosporidium felis and C. meleagridis in persons with HIV, Portugal. Emerg. Infect. Dis. 10: 2256–2257. McGlade, T.R., Robertson, I.D., Elliot, A.D., Read, C. and Thompson, R.C.A. 2003. Gastrointestinal parasites of domestic cats in Perth, Western Australia. Vet. Parasitol. 117: 251–262. McKenzie, D.M. and Diffay, B.C. 2000. Diarrhoea associated with cryptosporidial oocyst shedding in a quarterhorse stallion. Aust. Vet. J. 78: 27–28. McLauchlin, J., Amar, C., Pedraza-Díaz, S. and Nichols, G.L. 2000. Molecular epidemiological analysis of Cryptosporidium spp. in the United Kingdom: results of genotyping Cryptosporidium spp. in 1705 fecal samples from humans and 105 fecal samples from livestock animals. J. Clin. Microbiol. 38: 3984–3990. McReynolds, C.A., Lappin, M.R., Ungar, B., McReynolds, L.M., Bruns, C., Spilker, M.M., Thrall, M.A. and Reif, J.S. 1999. Regional seroprevalence of Cryptosporidium parvum-specific IgG of cats in the United States. Vet. Parasitol. 80: 187–195. Miller, D.L., Liggett, A., Radi, Z.A. and Branch, L.O. 2003. Gastrointestinal cryptosporidiosis in a puppy. Vet. Parasitol. 115: 199–204. Milstein, T.C. and Goldsmid, J.M. 1995. The presence of Giardia and other zoonotic parasites of urban dogs in Hobart, Tasmania. Aust. Vet. J. 72: 154–155. Molbak, K., Aaby, P., Hojlyng, N. and da Silva, A.P. 1994. Risk-factor for Cryptosporidium diarrhea in early childhood: a case-control study from Guinea-Bissau, West Africa. Am. J. Epidemiol. 139: 734–740. Monticello, T.M., Levy, M.G., Bunch, S.E. and Fairly, R.A. 1987. Cryptosporidiosis in a feline leukemia viruspositive cat. J. Am. Vet. Med. Assoc. 191: 705–706. Morgan, U.M., Sargent, K.D., Elliot, A. and Thompson, R.C.A. 1998. Cryptosporidium in cats additional evidence for C. felis. Vet. J. 156: 159–161. Morgan, U., Weber, R., Xiao, L., Sulaiman, I., Thompson, R.C.A., Ndiritu, W., Lal, A., Moore, A. and Desplazes, P. 2000a. Molecular characterization of Cryptosporidium isolates obtained from human immunodeficiency virus-infected individuals living in Switzerland, Kenya, and the United States. J. Clin. Microbiol. 38: 1180–1183. Morgan, U.M., Xiao, L., Monis, P., Irwin, P.J., Fayer, R., Fall, A., Denholm, K.M., Limor, J., Lal, A.A. and Thompson, R.C.A. 2000b. Cryptosporidium in domestic dogs: the dog genotype. Appl. Environ. Microbiol. 66: 2220–2223. Mtambo, M.M., Nash, A.S., Blewett, D.A., Smith, H.V. and Wright, S. 1991. Cryptosporidium infection in cats: prevalence of infection in domestic and feral cats in the Glasgow area. Vet. Rec. 129: 502–504.
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Mtambo, M.M.A., Nash, A.S., Wright, S.E., Smith, H.V., Blewett, D.A. and Jarrett, O. 1995. Prevalence of specific anti-Cryptosporidium IgG, IgM, and IgA in cat sera using an indirect immunofluorescence antibody test. Vet. Rec. 60: 37–43. Mtambo, M.M.A., Wright, S. E., Nash, A.S. and Blewett, D.A. 1996. Infectivity of a Cryptosporidium species isolated from a domestic cat (Felis domestica) in lambs and mice. Res. Vet. Sci. 60: 61–63. Mundim, M.J.S., Rosa, L.A.G., Hortncio, S.M., Faria, E.S.M., Rodrigues, R.M. and Cury, M.C. 2006. Prevalence of Giardia duodenalis and Cryptosporidium spp. in dogs from different living conditions in Uberlândia, Brazil. Vet. Parasitol. 144: 356–359. Nagy, B. 1995. Epidemiologic data on Cryptosporidium parvum infection of mammalian domestic animals in Hungary. Magyar Allatorvosok Lapja 50: 139–144. Nash, A.S., Mtambo, M.M.A. and Gibbs, H.A. 1993. Cryptosporidium infection in farm cats in the Glasgow area. Vet. Rec. 133: 576–577. Netherwood, T., Wood, J.L., Townsend, H.G., Mumford, J.A. and Chanter, N. 1996. Foal diarrhea between 1991 and 1994 in the United Kingdom associated with Clostridium perfringens, rotavirus, Strongyloides westeri and Cryptosporidium spp. Epidemiol. Infect. 117: 375–383. Nutter, F.B., Dubey, J.P., Levine, J.F., Breitschwerdt, E.B., Ford, R.B. and Stoskopf, M.K. 2004. Seroprevalences of antibodies against Bartonella henselae and Toxoplasma gondii and fecal shedding of Cryptosporidium spp, Giardia spp, and Toxocara cati in feral and pet domestic cats. J. Am. Vet. Med. Assoc. 225: 1394–1398. Olson, M.E., Thorlakson, C.L., Deselliers, L., Morck, D.W. and McAllister, T.A. 1997. Giardia and Cryptosporidium in Canadian farm animals. Vet. Parasitol. 68: 375–381. Pavlasek, I. and Ryan, U. 2007. The first finding of natural infection of Cryptosporidium muris in a cat. Vet. Parasitol. 144: 349–352. Pedraza-Diaz, S., Amar, C., Iversen, A.M., Stanley, P.J. and McLauchlin, J. 2001. Unusual Cryptosporidium species recovered from human feces: first description of C. felis and Cryptosporidium dog genotype from patients in England. J. Med. Microbiol. 50: 293–296. Pieniazek, N.J., Bornay-Llinares, F.J., Slemenda, S.B., da Silva, A.J., Moura, I.N.S., Arrowood, M.J., Ditrich, O. and Addiss, D.G. 1999. New Cryptosporidium genotypes in HIV-infected persons. Emerg. Infect. Dis. 5: 444–449. Pohjola, S. 1984. Survey of cryptosporidiosis in feces of normal healthy dogs. Nord. Vet.-Med. 36: 189–190. Ponce de León, P., Zdero, M. and Nocito, I. 1994. Investigación de Cryptosporidium sp. En heces de perro, en la ciudad de Rosario, Argentina. Acta Bioq. Clín. Latinoam. 28: 575–578. Reinemeyer, C.R., Kline, R.C. and Stauffer, G.D. 1984. Absence of Cryptosporidium oocysts in faeces of neonatal foals. Equine Vet. J. 16: 217–218. Ryan, U., Xiao, L., Read, C., Zhou, L., Lal, A.A. and Pavlasek, I. 2003. Identification of novel Cryptosporidium genotypes from the Czech Republic. Appl. Environ. Microbiol. 69: 4302–4307. Santín, M., Trout, J.M., Cortés Vecino, J.A., Dubey, J.P. and Fayer, R. 2006. Cryptosporidium, Giardia and Enterocytozoon bieneusi in cats from Bogota (Colombia) and genotyping of isolates. Vet. Parasitol. 141: 334–339. Sargent, K.D., Morgan, U.M., Elliott, A. and Thompson, R.C.A. 1998. Morphological and genetic characterisation of Cryptosporidium oocysts from domestic cats. Vet. Parasitol. 77: 221–227. Satoh, M., Matsubara-Nihei, Y., Sasaki, T. and Nakai, Y. 2006. Characterization of Cryptosporidium canis isolated in Japan. Parasitol. Res. Scorza, A.V., Brewer, M.M. and Lappin, M.R. 2003. Polymerase chain reaction for the detection of Cryptosporidium spp. in cat feces. J. Parasitol. 89: 423–426. Simpson, J.W., Burnie, A.G., Miles, R.S., Scott, J.L. and Lindsay, D.I. 1988. Prevalence of Giardia and Cryptosporidium infection in dogs in Edinburgh. Vet. Rec. 123: 445. Snyder, S.P., England, J.J., and McChesney, A.E. 1978. Cryptosporidiosis in immunodeficient Arabian foals. Vet. Pathol. 15: 12–17. Spain, C.V., Scarlett, J.M., Wade, S.E. and McDonough, P. 2001. Prevalence of enteric zoonotic agents in cats less than 1 year old in central New York State. J. Vet. Intern. Med. 15: 33–38. Sturdee, A.P., Bodley-Tickell, A.T., Archer, A. and Chalmers, R.M. 2003. Long-term study of Cryptosporidium prevalence on a lowland farm in the United Kingdom. Vet. Parasitol. 116: 97–113. Svobodova, V. Konvalinova, J. and Svobodova, M. 1994. Coprological and serological findings in dogs and cats with giardiosis and cryptosporidiosis. Acta Vet. Brno 63: 257–262.
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Turnwald, G.H., Barta, O., Taylor, H.W., Kreeger, J., Coleman, S.U. and Pourciau, S.S. 1988. Cryptosporidiosis associated with immunosuppression attributable to distemper in a pup. J. Am. Vet. Med. Assoc. 192: 79–81. Tzipori, S. and Campbell, I. 1981. Prevalence of Cryptosporidium antibodies in 10 animal species. J. Clin. Microbiol. 14: 455–456. Uga, S., Matsumura, T., Ishibashi, K., Yoda, Y., Yatomi, K. and Ataoka, N. 1989. Cryptosporidiosis in dogs and cats in Hyogo Prefecture, Japan. Jpn. J. Parasitol. 38: 139–143. Willard, M.D. and Bouley, D. 1999. Cryptosporidiosis, coccidiosis, and total colonic mucosal collapse in an immunosuppressed puppy. J. Am. Anim. Hosp. Assoc. 35: 405–409. Wilson, R.B., Holscher, M.A. and Lyle, S.J. 1983. Cryptosporidiosis in a pup. J. Am. Vet. Med. Assoc. 183: 1005–1006. Xiao, L. Bern, C., Limor, J., Sulaiman, I., Roberts, J., Checkley, W., Cabrera, L., Gilman, R.H. and Lal, A.A. 2001. Identification of 5 types of Cryptosporidium parasites in children in Lima, Peru. J. Infect. Dis. 183: 492–497. Xiao, L., Fayer, R., Ryan, U. and Upton, S.J. 2004. Cryptosporidium taxonomy: recent advances and implications for public health. Clin. Microbiol. Rev. 17: 72–97. Xiao, L. and Herd, R.P. 1994a. Epidemiology of equine Cryptosporidium and Giardia infections. Equine Vet. J. 26: 14–17. Xiao, L. and Herd, R.P. 1994b. Review of equine Cryptosporidium infection. Equine Vet. J. 26: 9–13. Xiao, L., Morgan, U.M. Limor, J., Escalante, A., Arrowood, M., Shulaw, W., Thompson, R.C., Fayer, R. and Lal, A.A. 1999. Genetic diversity within Cryptosporidium parvum and related Cryptosporidium species. Appl. Environ. Microbiol. 65: 3386–3391.
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18 Livestock
Mónica Santín and James M. Trout
CONTENTS I. II.
Introduction .................................................................................................................................. 451 Cryptosporidiosis in Cattle........................................................................................................... 452 A. Background ..................................................................................................................... 452 B. Clinical Features and Pathology ..................................................................................... 452 C. Detection and Identification............................................................................................ 454 D. Species and Genotypes ................................................................................................... 454 E. Prevalence........................................................................................................................ 455 F. Subtyping Cryptosporidium parvum .............................................................................. 458 III. Cryptosporidiosis in Sheep .......................................................................................................... 462 IV. Cryptosporidiosis in Goats........................................................................................................... 465 V. Cryptosporidiosis in Pigs ............................................................................................................. 466 VI. Cryptosporidiosis in Other Livestock .......................................................................................... 471 References........................................................................................................................... 472
I.
Introduction
For thousands of years humans have relied on domesticated livestock for food and fiber production. In many industrialized nations, livestock are often raised under intensive, high-density conditions, whereas other nations tend to use more traditional husbandry methods. Cryptosporidium infections have been reported in livestock worldwide, regardless of the husbandry system. Cryptosporidiosis, especially in young animals, can cause severe illness or death, resulting in decreased performance and production losses (de Graaf et al., 1999). A better understanding of Cryptosporidium species in animals and transmission potential can lead to better control strategies, resulting in healthier animals and greater production efficiency, which are essential to meet the ever-growing demand for food and fiber products. From the perspective of human health, many livestock harbor Cryptosporidium species/genotypes infectious for humans. The close association of humans and livestock, as well as the ability of runoff from animal operations to contaminate water supplies, presents a risk of human infection. Even with modern medical treatment, cryptosporidiosis can be debilitating to healthy individuals and can result in significant morbidity and mortality in special populations such as children, the elderly, and the immunocompromised. Treatment of cryptosporidiosis in both humans and animals remains problematic. A wide variety of compounds have been tested, although few have been found to be effective; likewise, attempts at developing vaccines have met with mixed success (See Chapter 9; Jenkins, 2004; Smith and Corcoran, 2004) . Studies in animals suggest that hyperimmune bovine colostrum, paromomycin, and nitazoxanide are effective in treating Cryptosporidium infections (Fayer and Ellis, 1993; Tzipori et al., 1994; Theodos et al., 1998; Jenkins et al., 1999; Perryman et al., 1999). However, treatment regimens and associated
451
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expenses make it unlikely that these compounds would be used in the livestock industry, limiting treatment to supportive therapy such as fluid and electrolyte replacement. There has been much progress in defining the Cryptosporidium species and genotypes present in livestock, their host ranges, and their transmission dynamics, especially for species/genotypes that cause disease in animals or that are transmissible to humans. Application of this information can improve the ability to control the spread of cryptosporidial infections, resulting in significant benefits to both animal and human health.
II. A.
Cryptosporidiosis in Cattle Background
Years of prevalence data have documented the worldwide occurrence of cryptosporidiosis in cattle. Point prevalence studies, in which a single sample is collected, indicate a wide range in the number of infected cattle, whereas longitudinal studies, in which the same animals are repeatedly sampled over time, often show a cumulative prevalence of 100%. Infections in young animals are frequently associated with morbidity, and sometimes mortality, and these young animals are known to harbor zoonotic species, representing a potential public health risk. Bovine cryptosporidiosis was first reported over three decades ago when histological examination of the jejunum of an 8-month-old heifer with chronic diarrhea revealed stages of the parasite (Panciera et al., 1971). Subsequently, cryptosporidial infections were reported in younger calves with diarrhea from both dairy and beef herds (Meuten et al., 1974; Morin et al., 1976). Initially, based on microscopic identification of oocysts, cattle were reported to harbor two species, C. parvum and C. muris, infecting the intestine and the abomasum, respectively (Panciera et al., 1971; Anderson, 1987). After application of molecular techniques and cross-species transmission studies, C. muris from cattle was properly identified as a new species, C. andersoni (Lindsay et al., 2000). Identification, prior to the use of molecular analysis, was based on oocyst size (C. parvum, 4.5 × 5.5 µm and C. andersoni, 5.5 × 7.4 µm) and/or the location of the infection in the intestine or abomasum for C. parvum and C. andersoni, respectively (Sunnotel et al., 2006). Cryptosporidium parvum was generally associated with diarrhea in young calves, whereas C. andersoni was identified in asymptomatic adults. Molecular analysis techniques have identified additional Cryptosporidium species and genotypes in cattle (Table 18.1) and are discussed in Section D.
B.
Clinical Features and Pathology
Cryptosporidium parvum infections in calves are often characterized by profuse watery diarrhea with acute onset. Diarrhea is accompanied by the shedding of high numbers of oocysts, up to 107 oocysts per gram of feces (Fayer et al., 1998); a single calf can potentially produce 1010 oocysts over the course of an infection (Nydam et al., 2001). Numerous studies have shown a significant correlation between C. parvum infection and diarrhea (Fayer et al., 1998; Adesiyun et al., 2001; Bjorkman et al., 2003; TrotzWilliams et al., 2005; Singh et al., 2006; Roy et al., 2006). Infection can also be accompanied by depression, weakness, and anorexia (Howerth, 1981; Tzipori et al., 1983). In experimental infections in calves, diarrhea occurred as early as 3 days postinfection, and lasted 4 to16 days; intensity (diarrhea and oocyst production) varied markedly between individuals (Fayer et al., 1998). Under natural conditions, calves appear to acquire infections shortly after birth; the highest rates are reported in calves 1 to 3 weeks of age (Huetink et al., 2001; Nydam et al., 2001; Castro-Hermida et al., 2002b; Santín et al., 2004), with oocyst shedding lasting an average of 12 days and diarrhea lasting an average of 8 days (Castro-Hermida et al., 2002b). The majority of C. parvum infections appear to be limited to calves under 8 weeks of age (Santín et al., 2004; Fayer et al., 2006, 2007; Langkjær et al., 2007). Parasites can be found in the epithelium of the small and large intestines (Tzipori et al., 1983; Heine et al., 1984b), but appear most numerous in the lower small intestine (Heine et al., 1984b). Enteritis, decreased villous length, villous atrophy, and fusion of villi have been observed in the small intestine (Howerth, 1981;
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TABLE 18.1 Species and Genotypes of Cryptosporidium Reported in Cattle Cryptosporidium Species/Genotypes
Frequency
Geographic Distribution
C. parvum
Common
Worldwide
C. bovis
Common
Worldwide
C. andersoni
Common
Worldwide
Deer-like genotype
Common
Worldwide
C. suis
Sporadic
United States and Zambia
C. suis-like Pig genotype II C. hominis
Sporadic Sporadic Sporadic
Demark Demark Scotland and Korea
C. felis C. canis
Rare Experimental infection only
Poland United States
Selected References Fayer et al., 2000 Peng et al., 2003 Becher et al., 2004 Santín et al., 2004 Langkjær et al., 2007 Starkey et al., 2006b Santín et al., 2004 Fayer et al., 2006, 2007 Feng et al., 2007 Geurden et al., 2006 Langkjær et al., 2007 Santín et al., 2004 Fayer et al., 2006, 2007 Feng et al., 2007 Geurden et al., 2006 Langkjær et al., 2007 Santín et al., 2004 Fayer et al., 2006, 2007 Feng et al., 2007 Geurden et al., 2006 Langkjær et al., 2007 Fayer et al., 2006 Geurden et al., 2006 Langkjær et al., 2007 Langkjær et al., 2007 Smith el al., 2005 Park et al., 2006 Bornay-Llinares et al., 1999 Fayer et al., 2001
Tzipori et al., 1983; Heine et al., 1984b); no lesions have been seen in the large intestine (Tzipori et al., 1983). Scanning and transmission electron microscopy of intestinal tissue revealed intracellular development, with trophozoites and meronts in the apical region of epithelial cells in the brush border; merozoites were also seen free in the lumen and in the process of invading epithelial cells (Pearson and Logan, 1983). Hypercellularity of the lamina propria and hyperplastic crypt epithelium are also associated with infection in the small intestine (Howerth, 1981; Heine et al., 1984b). Uninfected young calves typically have low percentages of CD4+ and CD8+ lymphocytes in the intraepithelial and lamina propria regions, although following infection, total intraepithelial lymphocytes and the percentages of CD4+ and CD8+ cells increase (Ruest et al., 1997; Fayer et al., 1998). These Tlymphocyte subsets have been reported to control cryptosporidial infections in mice (Ungar et al., 1991; Chen et al., 1993; McDonald et al., 1994); thus, the relative lack of these cells in young calves could explain their high susceptibility to C. parvum infection. Although preweaned calves are most susceptible to infection, not all C. parvum isolates exhibit similar virulence in terms of diarrhea severity and oocyst production. Such variation among isolates has been demonstrated in human infectivity studies and in calves using human-derived Cryptosporidium (Pozio et al., 1992; Okhuysen et al., 1999). Experimental calf infections with 5 × 106 oocysts isolated from calves in Maryland did not routinely induce clinical cryptosporidiosis (Fayer et al., 1985). Subsequently, infections with 1 × 104 oocysts isolated from calves in Alabama (AUCP-1 or Auburn-1 isolate) induced severe clinical disease in experimentally infected calves (Fayer and Ungar, 1986). Interestingly, after being passaged through calves for years at the ARS research facility, Beltsville, MD, the AUCP-1 strain began to lose pathogenecity. Over a period of 6 to
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12 months (serial passage through 6 to 7 calves), severity of clinical signs and oocysts output steadily decreased until inoculation with 1 × 106 oocysts resulted in no diarrhea and fewer than 106 total oocysts were recovered from a calf (R. Fayer and J. Trout, unpublished observations); initially, infection with AUCP-1 resulted in oocyst production rates of 107/g of feces (Fayer et al., 1998). Similar events occurred with the AUCP-1 isolate at Colorado State University (J. Vetterling, personal communication) and with the KSU isolate at Kansas State University (S. Upton, personal communication). During this same period, AUCP-1 was serially passaged through 40 groups of immunosuppressed C57BL/6 mice (4 mice/group), at Beltsville. Although the mice showed no clinical signs, oocyst output was consistently high; when oocysts isolated from the mice were inoculated into a calf, the AUCP-1 isolate exhibited typical virulence (R. Fayer and J. Trout, unpublished observations). Thus, it appears that not only do naturally occurring isolates of C. parvum vary in their level of virulence, but also that pathogenecity can change over time. It is not known what factors led to the observed loss of virulence in serial passage through calves or why virulence was unaffected by passage through mice. Cryptosporidium andersoni infects the gastric glands of the abomasum (Anderson, 1987, 1990) and has been reported in calves older than 4 weeks of age (Olson et al., 1997a; Santín et al., 2004; Kvac et al., 2006). Infections produce no diarrhea and follow a more chronic course than that of C. parvum; oocyst production is less than that of C. parvum infections (Kvac and Vitovec, 2003). Beef cattle shed C. andersoni oocysts from arrival at a feedlot through slaughter (Anderson, 1998), and dairy cows continuously shed oocysts for over a year (Esteban and Anderson, 1995). Although there are no visible clinical signs of infection, upon necropsy the gastric glands were dilated, the gastric mucosa was pale and thickened, and mucosal hyperplasia and inflammatory cell infiltration into the lamina propria of infected areas were evident (Anderson, 1998; Kvac and Vitovec, 2003; Masuno et al., 2006). Plasma pepsinogen was elevated (Anderson, 1998). Some infected beef cattle evaluated for weight gain performed well, others performed poorly (Anderson, 1998). Cryptosporidium andersoni-infected dairy cows reportedly produced significantly less milk than uninfected herd mates (Esteban and Anderson, 1995). Cattle infected with other Cryptosporidium species and genotypes (Table 18.1) (discussed in the following text) exhibited no overt clinical signs, there is no histological information, and there are no reports of subclinical pathology.
C.
Detection and Identification
Detection and identification protocols are sometimes used directly on fecal specimens. Frequently, however, feces are subjected to methods that concentrate oocysts and remove debris to increase the sensitivity of detection and/or remove potential inhibitors of PCR. Many of these techniques have been reviewed and the advantages and disadvantages of each method discussed (Chapter 6; Zarlenga and Trout, 2004; Sunnotel et al., 2006). For decades, microscopy was the sole method for detecting oocysts. At first, direct smears were routinely stained using a modified acid-fast protocol. Later, immunofluorescent antibody techniques became the method of choice. Microscopic identification was tedious, required a highly trained eye, sensitivity was generally low, and unless oocysts differed significantly in size, species identification was impossible. Serologic assays to detect antibodies, such as dot blot, Western blot, and ELISA indicated only past exposure to Cryptosporidium and generally could not identify species or genotypes. Fecal antigen-based tests, often in ELISA or dot blot formats, could identify current infections, but not species or genotypes. Immunofluorescent antibody stain (IFA) was found to be more sensitive than acid-fast staining (Quílez et al., 1996b), and PCR was found to be more sensitive than IFA (Santín et al., 2004). PCR has provided highly sensitive detection of infections that would be missed by other methods. Improved sensitivity has been demonstrated in detecting oocysts from both cats and postweaned calves (Scorza et al., 2003; McGlade et al., 2003; Santín et al., 2004).
D.
Species and Genotypes
Although several genes can be used to differentiate species and genotypes of Cryptosporidium, the most frequently used gene for species and genotype identification has been the SSU-rRNA gene (aka the 18S
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gene). Molecular analysis of Cryptosporidium isolated from cattle has resulted in the identification of seven species and three genotypes: C. parvum, C. andersoni, C. bovis, C. canis, C. felis, C. hominis, C. suis, C. suis-like genotype, Cryptosporidium deer-like genotype, and Cryptosporidium pig genotype II (Bornay-Llinares et al., 1999; Fayer et al., 2001; Santín et al., 2004; Smith et al., 2005; Fayer et al., 2006; Feng et al., 2007; Geurden et al., 2006; Langkjær et al., 2007; Park et al., 2006; Starkey et al., 2006a) (Table 18.1). In the few large-scale prevalence studies in cattle to date, C. parvum, C. andersoni, C. bovis, and Cryptosporidium deer-like genotype have been found most frequently (Xiao et al., 2004; Santín et al., 2004; Fayer et al., 2006; Feng et al., 2007; Geurden et al., 2006; Langkjær et al., 2007). In contrast, C. hominis has been found in only a few cattle in Scotland, India, and Korea (Smith el al., 2005; Park et al., 2006; Feng et al., 2007); C. suis was found in a calf in the United States and a calf in Zambia (Fayer et al., 2006; Geurden et al., 2006); C. suis-like genotype was found in three cattle in Denmark (Langkjær et al., 2007); Cryptosporidium pig genotype II was found in a cow in Denmark (Langkjær et al., 2007); and C. felis was found in one cow in Poland (Bornay-Llinares et al., 1999). Although C. canis infected calves experimentally (Fayer et al., 2001), natural infections have not been detected. It is unclear if the less frequently detected species and genotypes are maintained within cattle populations or result from occasional contact with the primary host.
E.
Prevalence
Since the first report of Cryptosporidium in a calf, there have been many prevalence studies based on numerous detection and identification techniques, with different levels of sensitivity resulting, in part, in considerable variation in reports on the prevalence of bovine cryptosporidiosis. Most surveys measure point prevalence, in which a single sample is collected per animal in a given herd (Olson et al., 1997b; Atwill et al., 1999; Castro-Hermida et al., 2002a; Enemark et al., 2002). A smaller number of longitudinal surveys detect the cumulative prevalence over time from multiple samples collected from the same animal (Xiao and Herd, 1994; O’Handley et al., 1999; Uga et al., 2000; CastroHermida et al., 2002b). Point prevalence studies tend to underestimate the true prevalence in a population, because the patent period can be missed and oocyst excretion can be intermittent; however, larger numbers of animals can be examined than in longitudinal studies, which involve greater time, financial, and logistical efforts. Based on its large oocyst size, only C. andersoni can be identified with certainty in cattle feces by microscopy. Using molecular analysis, six species of Cryptosporidium and at least three genotypes, with oocysts of similar size and morphology, can be detected in cattle feces. It is therefore essential that molecular techniques be incorporated into prevalence studies. Studies that screen fecal samples by IFA and subject only the positive samples to molecular analysis can skew results in favor of species or genotypes that produce higher number of oocysts. Accurate prevalence data require uniform application of molecular analysis to all fecal samples from a given population. Cattle are the primary nonhuman species impacted by cryptosporidiosis. Thus, more prevalence studies have been conducted on cattle than on any other animals. Prevalence studies prior to the mid-1990s have been reviewed (Angus, 1990; Casemore et al., 1997; Fayer et al., 1997). Subsequent prevalence data based on microscopic analysis underscore the worldwide presence of Cryptosporidium in cattle. In a multitude of surveys, the reported prevalence in cattle ranged from 0 to 100% (Tables 18.2–18.4). Typically, a higher prevalence was seen in younger animals compared to older ones (Becher et al., 2004; Castro-Hermida et al., 2006; Maddox-Hyttel et al., 2006; Roy et al., 2006; Starkey et al., 2006b) (Figure 18.1). Based on oocyst size, C. parvum-like oocysts were most often seen in young calves, and C. andersoni was found more frequently in older cattle, (Kvac et al., 2006). Studies on beef cattle suggest a periparturient rise in excretion of C. andersoni oocysts (Ralston et al., 2003), but not of C. parvumlike oocysts (Atwill et al., 2003). In dairy cattle, C. parvum-like oocysts were not detected in periparturient cows in two studies (Atwill et al., 1998; Atwill and Pereira, 2003), but were detected in a third survey (Castro-Hermida et al., 2002a). However, without molecular analysis the actual species or genotypes remain unknown. Studies using molecular analysis to identify Cryptosporidium species and genotypes are presented in Tables 18.5 to 18.7. A series of large-scale, point prevalence studies examined dairy cattle in the eastern
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TABLE 18.2 Prevalence of Cryptosporidium in Dairy Cattle Determined by Nonmolecular Detection Methods, Arranged by Country
Location Australia Australia Brazil
Number of Animals/Number of Farms
Age
Prevalence (% positive) a
Canada Canada Canada Canada Canada Czech Republic
54/2 40/2 30 cows 30 calves 3 farms 386/20 505 farms 20/1 500/51 30/1 4273/11
Denmark Denmark England England France France India Japan Korea Mexico
255/50 895/50 367/NS 516/NS 382/6 1628/7f 90/4 30/1 84/NS 512/31
Lactating cows 0–12 m < 6–8 w Adult 8–15 d 4–12 d Varied 0–30 d NS 1–30 d
15 88.7c 100a 40.6 0a Preweaned 27.1; 0.9b Postweaned 0; 32.5b 17 40 52 3.5d 51.8e 17.9 40 93a 100 25
New Zealand Norway Spain Spain Sweden Tanzania Trinidad Venezuela
185/24 1386/136 535/12 734/60 270/75 486/5 333/NS 75/4
Neonates 3–183 d 21 d Varied 0–90 d Varied Adults 2–20 w
21.2 12 47.5 14.2 8.1 5.3 3.3 29.3
United States—Maryland
43/2
Adult
United States—Maryland United States—New York United States—New York
23/1 557/11 2943/109
2–12 m 0–12 w Varied
United States—New York United States—New York
6530/39 453/19
Varied Varied
Zambia
250
1–90 d
4.7 7b 8.7 8.3 0.9 1.1b 11.4 Overall—3.2 < 60 d of age—20 42.8g
Reference
0–12 w Adult Adults and calves 0–14 m
48.1 0 Adults—0 Calves—47a,b
Becher et al., 2004 Becher et al., 2004 Pena et al., 1997
0–24 w Calves 0–120 d 7–21 d Adult Pre- and postweaned
Olson et al., 1997a Ruest et al., 1998 O’Handley et al., 1999 Trotz-Williams et al., 2005 Uehlingher et al., 2006 Kvac et al., 2006
Note: d = day; w = week; m = month; NS = not specified. a Longitudinal design, cumulative prevalence. b Reported as C. andersoni (formerly known as C. muris—bovine) based on oocyst size. c Percentage of farms where Cryptosporidium was detected. d Did not differentiate Cryptosporidium parvum-like from C. andersoni. e Highest prevalence of multiple sampling days. f Administrative regions of France. g For molecular data on selected samples, see Tables 18.5–18.7.
Maddox-Hyttel et al., 2006 Maddox-Hyttel et al., 2006 Sturdee et al., 2003 Sturdee et al., 2003 Naciri et al., 1999 Lefay et al., 2000 Singh et al., 2006 Uga et al., 2000 Yu et al., 2004 Maldonado-Camargo et al., 1998 Grinberg et al, 2005 Hamnes et al., 2006 Castro-Hermida et al., 2002a Castro-Hermida et al., 2006 Bjorkman et al., 2003 Mtambo et al., 1997 Adesiyun et al., 1996 Surumay-Vilchez and Alfaro, 2000 Fayer et al., 2000 Fayer et al., 2000 Sischo et al., 2000 Wade et al., 2000 Starkey et al., 2005 Starkey et al., 2006a Geurden et al., 2006
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TABLE 18.3 Prevalence of Cryptosporidium in Beef Cattle Determined by Nonmolecular Detection Methods, Arranged by Country
Location
Number of Animals/Number of Farms
Canada
20/1
Canada
669/39
Canada Canada Canada Czech Republic
193/10 560/59 605/100 2748/11
England France Korea Spain United States—California United States—California
220/NS 3080/94 863/NS 309/10 1321/38 120/3
United States—Maryland Zambia
118/1 238
Age 0–6 m
Prevalence (% positive)
5 0a 2–14 y 18.4 10.6a 2–70 d 12.6 Adults 1.1 0–211 d 3.1 Pre- and postweaned Preweaned 0.8; 0.3a Postweaned 0; 21.2a Adult 3.6 0–30 d 15.6 NS 20.2 ≤ 21 d 48.5 Varied 3.9 Adult prepart—8.3 Pre- and postpart postpart—5.8 7–9 m 28.8 2–70 d 87
Reference Ralston et al., 2003 McAllister et al., 2005 McAllister et al., 2005 Gow and Waldner, 2006 Gow and Waldner, 2006 Kvac et al., 2006
Sturdee et al., 2003 Bendali et al., 1999 Yu et al., 2004 Castro-Hermida et al., 2002a Atwill et al., 1999 Atwill et al., 2003 Fayer et al., 2000 Geurden et al., 2006
Note: d = day; m = month; y = year; NS = not specified. a b
Reported as C. andersoni based on oocyst size. For molecular data on selected samples, see Tables 18.5–18.7.
United States (Santín et al., 2004; Fayer et al., 2006, 2007); data are summarized in Figure 18.2. Virtually all infections in calves 8 weeks of age and younger were caused by C. parvum, although low levels of C. bovis, Cryptosporidium deer-like genotype, and C. andersoni were found in some calves. The number of infections caused by C. parvum dropped dramatically in postweaned calves (2 to 11 months of age), whereas levels of C. bovis, Cryptosporidium deer-like genotype, and C. andersoni increased, with C. bovis the predominate species in this age group. In animals older than 12 months, C. andersoni was the most prevalent species. In Denmark, C. parvum was also found to be the primary species in calves 1 month of age and younger, and similar changes in species and genotypes were found in calves 3 to 12 months of age, with C. bovis the predominate species (Langkjær et al., 2007). Indeed, C. bovis and Cryptosporidium deer-like genotype appear to be geographically widespread, having been reported in cattle from China, Denmark, India, and various regions of the United States (Santín et al., 2004; Fayer et al., 2006, 2007; Feng et al., 2007; Langkjær et al., 2007; Starkey et al., 2006b). Approximately half the infections in adult cattle in the United States were C. andersoni (Fayer et al., 2006, 2007). In Denmark, the Czech Republic, and Japan, a higher prevalence of C. andersoni in older animals has been reported as well (Enemark et al., 2002; Kvac and Vitovec, 2003; Sakai et al., 2003; Kvac et al., 2006). Interestingly, Langkjær et al. (2007), also in Denmark, did not detect C. andersoni in animals of any age. The overall prevalence of Cryptosporidium declines with increasing age (Figure 18.1) (Santín et al., 2004; Fayer et al., 2006, 2007; Kvac et al., 2006; Langkjær et al., 2007). Therefore, the species and genotypes that constitute a large proportion of the infections in adult cattle are actually present in relatively few animals in the herd. As noted in Section D, several species and genotypes usually associated with other hosts have been infrequently reported in cattle. The sources are unknown.
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Cryptosporidium and Cryptosporidiosis, Second Edition TABLE 18.4 Prevalence of Cryptosporidium in Cattle Determined by Nonmolecular Detection Methods, Arranged by Country (Animal Type is Mixed Dairy and Beef, or Could Not be Determined from Reference)
Location
Number of Animals/Number of Farms
Austria Canada England France France Germany Germany India Iraq Japan Korea Malaysia Northern Ireland Poland Spain Spain
230/100 104/6 272 311/40 440/189 4060b 998/NS 101/NS 60/NS 512/slaughterhouse 201e/NS 96/7 779/193 75/9 554/30 101/1
1–42 d Varied <8w 4–10 d 4–21 d NS NS Adults and calves NS Adults <4m NS <1m 3–13 d Varied 2 d–24 m
Switzerland Taiwan Trinidad Turkey Uganda Zambia
386/67 460/27 59/12 109/NS 50/5 256
0–3 m Varied Varied Calves Pre- and postweaned 1–84 d
Age
Prevalence (% positive) 11.7 20 23.2 86a 43.4 21.5 3.5 10.9 20 4.7c,d 14.4 25 37.4 51 19.7 26 59 3.7 37.6f 6.8 35.8 38 6.3g
Reference Haschek et al., 2006 Olson et al., 1997b Sturdee et al., 2003 Naciri et al., 1999 Lefay et al., 2000 Joachim et al., 2003 Epe et al., 2004 Khubnani et al., 1997 Mahdi and Ali, 2002 Kaneta and Nakai, 1998 Wee et al., 1996 Farizawati et al., 2005 Thompson et al., 2006 Bednarska et al., 1998 Quílez et al., 1996d Ares-Mazas et al., 1999 Lentze et al., 1999 Watanabe et al., 2005 Adesiyun et al., 2001 Sahal et al., 2005 Nizeyi et al., 2002 Geurden et al., 2006
Note: d = day; w = week; m = month; NS = not specified. a b c d
e f g
F.
Highest prevalence of multiple sampling days. Samples submitted to diagnostic laboratories. Reported as C. andersoni based on oocyst size. Kaneta and Nakai (1998) reported these oocysts induced infection in the stomachs of mice, which is not consistent with C. andersoni. Only diarrheic animals were sampled. Two sizes of oocysts, corresponding to C. parvum and C. andersoni, noted. For molecular data on selected samples, see Tables 18.5–18.7.
Subtyping Cryptosporidium parvum
As Cryptosporidium species were subjected to increased molecular scrutiny, genetic subtype families as well as subtypes within families could be identified within C. parvum and C. hominis (Cacciò et al., 2000; Feng et al., 2000; Strong et al., 2000; Peng et al., 2001, 2003; Leav et al., 2002; Alves et al., 2003; Gasser et al., 2003; Tanriverdi et al., 2003; Wu et al., 2003; Xiao and Ryan, 2004). Because C. parvum is the zoonotic species most commonly identified in cattle, it has been the focus of subtyping analysis to determine its genetic diversity and to serve as a tool for source tracking (Chapter 5). Most subtyping tools have been based on the 60-kDa glycoprotein gene (GP60 gene). Sequence analysis of the gene has revealed several subtype families (alleles) and subtypes within C. parvum. Most cattle isolates belong to the C. parvum subtype family IIa (Table 18.8), the zoonotic subtype family of C. parvum found in humans and ruminants. The anthropozoonotic C. parvum subtype family IIc (previously known as Ic) has not been found in cattle. Within the IIa subtype family, the most common subtype found worldwide was IIaA15G2R1; e.g., Portugal (61 out of 72 isolates) (Alves et al., 2003, 2006), India (5 out of 9) (Feng et al., 2007), or in the United States (135 out of 175) (Xiao et al., 2007b) (see Table 18.8). In Northern Ireland, a greater genetic diversity within allele IIa was observed; the most
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TABLE 18.5 Identification of Cryptosporidium Species and Genotypes in Dairy Cattle, Using Molecular-Based Analysis Methods, Arranged by Country Location
Number of Samples
Age
Gene
Species/Genotype Prevalence
Australia
6 isolatesa
0–12 w
China
6 isolates
Denmark
154c
India
12 isolates
New Zealand
658
Pre- and SSU-rRNA 5 isolates were C. bovis postweaned 1 isolate was Cryptosporidium DLGb calves Varied SSU-rRNA 50.6% of animals + for C. parvum HSP-70 37.0% of animals + for C. bovis 7.8% of animals + for Cryptosporidium DLG 1.9%, 3 animals + for C. suis-like genotype 0.7%, 1 animal + for Cryptosporidium pig genotype II 1.9%, 3 animals, with atypical isolates Pre- and SSU-rRNA 11 isolates were C. parvum postweaned 1 isolate was C. bovis calves Cows/calves SSU-rRNA 5.3% of animals + for C. parvum
Northern Ireland 224
<1 m
United States— multiple states
393
5 d–2 m
United States— multiple states
447
3–11 m
United States— multiple states
571
1–2 y
United States— Michigan United States— New York
248
Cows/calves
115c
NS
United States— Georgia
32
Varied
SSU-rRNA All isolates were C. parvum
SSU-rRNA 95.1% of isolates were C. parvum 3.6% of isolates were C. bovis 1.3% of isolates were Cryptosporidium DLG SSU-rRNA 35.1% of animals + for C. parvum 3.6% of animals + for C. bovis 2.0% of animals + for Cryptosporidium DLG SSU-rRNA 0.2%, 1 animal + for C. parvum 14.5% of animals + for C. bovis 3.4% of animals + for C. andersoni 8.1% of animals + for Cryptosporidium DLG SSU-rRNA 0.7% of animals + for C. parvum 4.2% of animals + for C. bovis 5.1% of animals + for C. andersoni 1.8% of animals + for Cryptosporidium DLG 0.2%, 1 animal + for C. suis SSU-rRNA 22.6% of animals + for C. parvum 1.6% of animals + for C. andersoni SSU-rRNA 60.8% of isolates were C. parvum 36.5% of isolates were C. bovis 2.6% of isolates were Cryptosporidium DLG SSU-rRNA 6 isolates were C. parvum 15 isolates were C. bovis 6 isolates were Cryptosporidium DLG 1 isolate was mixed C. parvum/ C. andersoni 1 isolate mixed C. bovis/Cryptosporidium DLG
Reference Becher et al., 2004 Feng et al., 2007
Langkjær et al., 2007
Feng et al., 2007
Learmonth et al., 2003 Thompson et al., 2006
Santín et al., 2004
Santín et al., 2004
Fayer et al., 2006
Peng et al., 2003 Starkey et al., 2006b
Feng et al., 2007
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TABLE 18.5 (CONTINUED) Identification of Cryptosporidium Species and Genotypes in Dairy Cattle, Using Molecular-Based Analysis Methods, Arranged by Country Location Wales
Zambia
Number of Samples
Age
101 (ground- Varied collected samples)a 32 isolates 1–90 d
Gene
Species/Genotype Prevalence
Reference
SSU-rRNA 7/16 positive samples were C. andersoni Robinson et al., 2006 SSU-rRNA By SSU-rRNA: HSP-70 22 isolates were C. parvum 10 isolates were C. bovis By HSP-70: 21 isolates were C. parvum 9 isolates were C. bovis 2 did not amplify
Geurden et al., 2006
Note: d = day; w = week; m = month; y = year; NS = not specified; + = positive. a b c
Analyzed all, or a subset of, samples positive by microscopy. Cryptosporidium “deer-like genotype.” Did not report prevalence, only species/genotypes present in Cryptosporidium-positive samples.
TABLE 18.6 Identification of Cryptosporidium Species and Genotypes in Beef Cattle Using Molecular-Based Analysis Methods, Arranged by Country Location Czech Republic
Number of Samples
Age
Gene
Species/Genotype Prevalence
Reference
56.8% of samples + for C. andersoni 92.9% calves + for C. andersoni 43.8% of animals + for C. andersoni
Kvac and Vitovec, 2003 Kvac and Vitovec, 2003 Enemark et al., 2002
1–35 w
SSU-rRNA
Czech Republic
117 calves/599 samples 96
Adult
SSU-rRNA
Denmark
292
Varied
SSU-rRNA
Ireland
288
Adult
NS
United States— multiple states Zambia
5274a
Feedlot
SSU-rRNA
8 isolates
2–70 d
SSU-rRNA HSP-70
23.6% of animals + for C. andersoni 10.6% reported as C. parvum (by microscopy) 3.5% of animals + for C. parvum 3.8% of animals + for C. andersoni 4 samples were C. parvum By SSU-rRNA: 3 isolates were C. parvum 1 isolate was C. suis 4 did not amplify By HSP-70: 7 isolates were C. parvum 1 isolate was C. suis
Moriarty et al., 2005 Atwill et al., 2006 Geurden et al., 2006
Note: d = day; w = week; NS = not specified; + = positive a
Analyzed all, or a subset of, samples positive by microscopy.
common C. parvum subtype was IIaA18G3R1 (120 out of 214) followed by the subtype IIA15G2R1 (28 out of 214) (Thomson et al., 2006) (see Table 18.8). The high prevalence of subtype IIaA15G2R1 in calves worldwide and its frequent detection in humans in Australia, Japan, Kuwait, Northern Ireland, Portugal, Slovenia, and the United States (Strong et al., 2000; Glaberman et al., 2002; Abe et al., 2006; Alves et al., 2003, 2006; Peng et al., 2003; StanticPavlinic et al., 2003; Wu et al., 2003; Chalmers et al., 2005; Sulaiman et al., 2005) suggest that it is easily spread among cattle populations and readily transmitted to humans as well.
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TABLE 18.7 Identification of Cryptosporidium Species and Genotypes in Cattle, Using Molecular-Based Analysis Methods Arranged by Country (Animal Type is Mixed Dairy and Beef or Could Not be Determined from the Reference) Location
Number of Samples
Chile India Japan
127a 940a 480a
Varied 0–12 m 0–39 m
SSU-rRNA SSU-rRNA NS
Korea Scotland
17 411b
NS NS
Sweden
50
Calves
Taiwan Zambia
460/27 5 isolates
Varied 1–84 d
SSU-rRNA SSU-rRNA Microsatellite SSU-rRNA COWP Microsatellite SSU-rRNA SSU-rRNA HSP-70
Age
Species/Genotype Prevalence
Gene
Reference
1 sick adult female + for C. parvum 17.7% C. parvum 1.3% of animals + for C. parvum 3.3% of animals + for C. andersoni 35% of animals + for C. hominis 99.5% of animals + for C. parvum 0.5%, 2 animals, + for C. hominis Novel genotype of C. parvum
Neira-Otero et al., 2005 Roy et al., 2006 Sakai et al., 2003
1 isolate + for C. parvum By SSU-rRNA: 4 isolates were C. bovis 1 did not amplify By HSP-70: 5 isolates were C. bovis
Watanabe et al., 2005 Geurden et al., 2006
Park et al., 2006 Smith et al., 2005 Bjorkman and Mattsson, 2006
Note: d = day; m = month; NS = not specified; + = positive. a b
Analyzed all, or a subset of, samples positive by microscopy. From clinically ill animals.
50
40
30
20
10
0 < 2 mo
3-11 mo
12-24 mo
> 24 mo
Age FIGURE 18.1 Prevalence of Cryptosporidium against age in dairy cattle. (Adapted from Santín, M., Trout, J. M., Xiao, L., Zhou, L., Greiner, E. and Fayer, R. 2004. Prevalence and age-related variation of Cryptosporidium species and genotypes in dairy calves. Vet. Parasitol. 122: 103–117; Fayer, R., Santín, M, Trout, J.M. and Greiner, E. 2006. Prevalence of species and genotypes of Cryptosporidium found in 1- to 2-year-old dairy cattle in eastern United States. Vet. Parasitol. 135: 105112; Fayer, R., Santín, M, and Trout, J.M. 2007. Prevalence of Cryptosporidium species and genotypes in mature dairy cattle on farms in the eastern United States compared with younger cattle from the same locations. Vet. Parasitol. 145: 260–266.)
Knowledge regarding sources of environmentally detected oocysts and Cryptosporidium population structures within and between herds remains elusive. Subtyping holds promise as a valuable tool for tracking infection sources and examining transmission dynamics. However, despite geographic differences in distribution of specific alleles, too few isolates have been analyzed to understand the dynamics of C. parvum transmission among and between cattle and human populations.
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FIGURE 18.2 Percentage of each Cryptosporidium species or genotype represented in different age groups of dairy cattle. (Adapted from Santín, M., Trout, J. M., Xiao, L., Zhou, L., Greiner, E. and Fayer, R. 2004. Prevalence and age-related variation of Cryptosporidium species and genotypes in dairy calves. Vet. Parasitol. 122: 103–117; Fayer, R., Santín, M, Trout, J.M. and Greiner, E. 2006. Prevalence of species and genotypes of Cryptosporidium found in 1- to 2-year-old dairy cattle in eastern United States. Vet. Parasitol. 135: 105-112; Fayer, R., Santín, M, and Trout, J.M. 2007. Prevalence of Cryptosporidium species and genotypes in mature dairy cattle on farms in the eastern United States compared with younger cattle from the same locations. Vet. Parasitol. 145: 260–266.)
III. Cryptosporidiosis in Sheep Ovine cryptosporidiosis was first described in lambs with diarrhea in Australia by Baker and Carbonell (1974). The role of Cryptosporidium as a primary etiological agent of diarrhea in lambs was reported in the early 1980s in both natural and experimental infections (Angus et al., 1982; Snodgrass et al., 1984). Ovine cryptosporidiosis has subsequently been found worldwide (e.g., Xiao et al., 1993; Majewska et al., 2000; McLauchlin et al., 2000; Causapé et al., 2002; Alonso-Fresán et al., 2005; Ryan et al., 2005a; Santín et al., 2007). As one of the major enteropathogens associated with neonatal diarrhea, it causes significant morbidity and mortality in lambs (de Graaf et al., 1999). Economic losses reflect not only mortality but retarded growth rates after recovery, veterinary assistance, and increased labor costs associated with extra care. A wide range of Cryptosporidium prevalence in sheep has been reported, most based on microscopy (Table 18.9). PCR-based detection was demonstrated to be more sensitive than microscopy: in Australia, the prevalence was 2.6% by microscopy compared to 26.25% by PCR (Ryan et al., 2005a). In the United States Cryptosporidium was identified in 20.6% by microscopy compared to 50.8% by PCR (Santín et al., 2007). There is a close association between prevalence of infection and age of the animals, the highest prevalence found in lambs (Xiao et al., 1993; Abou-Eisha, 1994; Majewska et al., 2000; Causapé et al., 2002; Sturdee et al., 2003; Santín et al., 2007; Table 18.9). Of 583 lambs, 76.2% shed oocysts at 8 to 14 days of age; significantly higher rates of infection were observed in 1- to 21-day-old lambs (64.4%) than in 22- to 90-day-old lambs (23%) (Causapé et al., 2002). Similar results were obtained in Serbia; a higher prevalence was observed in lambs less than 30 days of age (45.3%) than in 30- to 90-day-old ˇ ´ et al., 2006). Cryptosporidium infection in lambs less than 30 days of age was lambs (38.7%) (Misic more likely to cause diarrhea than in older lambs, which were frequently asymptomatic. Apparently healthy adults and lambs can serve as a source of infection for other animals, possibly maintaining cryptosporidiosis within a flock between lambing periods. A periparturient rise in oocyst shedding was observed in ewes in Spain and in the United States (Xiao et al., 1994b; Ortega-Mora et al., 1999). In Spain, 13 of 14 ewes excreted oocysts at some time between 6 weeks before until 2 weeks after birth. The number of animals excreting oocysts increased 1 week before and 1 week after parturition.
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463 TABLE 18.8 Cryptosporidium Parvum GP60 Subtypes Identified in Cattle. Subtypes were Determined and Named Based on Both the Number of Trinucleotide Repeats and Mutations in the Nonrepeat Regions* Geographic Origin
Number of Samples
Canada United States Portugal
2 1 72
United States
33
Slovenia Italy Japan United Kingdom Japan United States India
4 1 1 2 3 6 9
Northern Ireland
214
Canada
36
GP60 Subtype of C. parvum (number of specimens) IIaA16G3R1 (2) IIaA18G5R1 (1) IIaA15G2R1 (61) IIaA16G2R1 (7) IIdA17G1 (4) IIaA15G2R2 (10) IIaA15G2R1 (8) IIaA16G2R1 (8) IIaA16G1R1 (3) IIaA16G3R1 (2) IIaA16G3R2 (2) IIaA17G1R1 (4)a IIaA13G2R1 (1) IIaA15G2R1 (1) IIaA20G3R1 (2) IIaA15G2R1 (3) IIaA15G2R1 (6) IIaA15G2R1 (5) IIaA13G2R2 (1) IIaA14G2R1a (1) IIaA14G2R1b (1) IdA15G1 (1)b IIaA18G3R1 (120) IIaA15G2R1 (28) IIaA17G2R1 (19) IIaA19G4R1 (15) IIaA20G3R1 (6) IIaA17G3R1 (5) IIaA19G3R1 (5) IIaA20G5R1 (3) IIaA18G2R1 (2) IIaA20G2R1 (2) IIaA16G3R1 (1) IIaA17G1R1 (1) IIaA18R1 (1) IIaA19G2R1 (1) IIaA20G4R1 (1) IIaA21G2R1 (1) IIa-unknown (5) New family (1) IIa15G2R1 (10) IIa16G2R1 (9) IIaA16G3R1 (8) IIaA16G1R1 (4) IIaA13G2R1 (2) IIaA17G2R1 (2) IIaA18G3R1 (1)
Reference Strong et al., 2000 Alves et al., 2003, 2006
Peng et al., 2003
Stantic-Pavlinic et al., 2003 Wu et al., 2003 Chalmers et al., 2005 Abe et al., 2006 Feng et al., 2006
Thompson et al., 2006
Trotz-Williams et al., 2006
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Cryptosporidium and Cryptosporidiosis, Second Edition TABLE 18.8 (CONTINUED) Cryptosporidium Parvum GP60 Subtypes Identified in Cattle. Subtypes were Determined and Named Based on Both the Number of Trinucleotide Repeats and Mutations in the Nonrepeat Regions* Geographic Origin United States (16 farms in 8 states)
* a b
Number of Samples 175
GP60 Subtype of C. parvum (number of specimens) IIaA15G2R1 (134) IIaA15G2R2 (10) IIaA11G2R1 (9) IIaA18G2R1 (9) IIaA17G2R1 (7) IIaA19G2R1 (3) IIaA11G2R1+IIaA15G2 R2 (1) IIaA11G2R1+IIaA15G2 R1 (1) IIaA18G2R1+IIaA19G2 R1 (1)
Reference Xiao et al., 2007
Nomenclature as established by Sulaiman et al., 2005. With a change of A to G after the trinucleotide repeats. C. hominis subtype.
TABLE 18.9 Prevalence of Cryptosporidium Reported from Sheep by Microscopy Country Australia Canada Egypt Egypt Hungary Iran Iraq Mexico Poland Serbia Spain Spain Trinidad and Tobago United Kingdom United Kingdom United States United States a
Number of Animals
Prevalence (% positive)
1647 sheep 89 sheep 120 lambs 100 lambs 83 ewes 53 lambs 215 sheep 45 sheep 556 lambs 664 ewes 60 lambs 99 ewes 126 lambs 583 lambs 205 ewes 69 lambs 90 lambs 90 lambs 255 lambs 250 ewes 32 lambs 23 ewes 31 lambs 32 ewes
2.6% 23% 36–82%a 24% in lambs 2.4% in ewes 22.6% 17.2 % 13.3% 33.5% in lambs 34% in ewes 18.3% in lambs 5% in ewes 42.1% 59% in lambs 7.8% in ewes 1.45% 20% 53–74%a 12.9% in lambs 6.4% in ewes 84.4% in lambs 17.4% in ewes 77.4% in lambs 25% in ewes
Varied with the detection method used.
Reference Ryan et al., 2005a Olson et al., 1997a Abd-El-Wahed, 1999 Abou-Eisha, 1994 Nagy, 1995 Nouri and Karami, 1991 Mahdi and Ali, 2002 Alonso-Fresán et al., 2005 Majewska et al., 2000 ˇ ´ et al., 2006 Misic Causapé et al., 2002 Villacorta et al., 1991 Kaminjolo et al., 1993 Chalmers et al., 2005 Sturdee et al., 2003 Xiao et al., 1993 Santín et al., 2007
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TABLE 18.10 Cryptosporidium Species/Genotypes Reported from Sheep With Molecular Data Cryptosporidium Species/Genotype (number of Isolates)
Country
C. parvum (1)
Spain
C. parvum (16) C. parvum (9) Novel sheep genotype (26) Cervine genotype (33) C. bovis (14) Pig genotype II (4) Marsupial genotype (4) C. suis (2) C. andersoni (1) C. hominis (1) Unique unknown genotype (1) Cervine genotype (48) C. bovis-like (7) C. parvum (2)
United Kingdom United Kingdom
Gene SSU-rRNA Acetyl-CoA synthetase COWP COWP
Reference Morgan et al., 1998 McLauchlin et al., 2000 Chalmers et al., 2002
Australia
SSU-rRNA HSP-70 (only in a subset of the isolates)
Ryan et al., 2005a
United States
SSU-rRNA
Santín et al., 2007
Lack of colostrum uptake, poor hygiene, and the presence other enteropathogens contribute to clinical disease. The risk of Cryptosporidium is higher in crowded flocks, probably because of hygienic and management conditions (Alonso-Fresán et al., 2005). Most cryptosporidial infections reported in sheep before genotyping were thought to be caused by C. parvum. More recently, other species and genotypes have been identified (Table 18.10). In the United Kingdom, a sheep genotype was the most common Cryptosporidium identified, being reported in 26 out of 35 samples examined (Chalmers et al., 2002); however, the identity of this genotype remains uncertain, as no molecular data were presented (Chalmers et al., 2002). In Australia, two genotypes were reported, a novel bovine genotype B (which was renamed as C. bovis by Fayer et al., 2005) and a previously unknown genotype were found in 14 and 1 of 60 samples examined, respectively (Ryan et al., 2005a). In the United States, a C. bovis-like genotype was found in 7 of 57 specimens (Santín et al., 2007). Two studies of Cryptosporidium isolates from sheep found that the cervine genotype, not C. parvum, was the most common Cryptosporidium in sheep (Ryan et al., 2005a; Santín et al., 2007). The cervine genotype has both a wide host and geographic range, having been found in sheep in Australia, lemur and white-tailed deer in North America, blesbok, mouflon, and nyala in the Czech Republic, and in humans in Canada, New Zealand, United Kingdom, Slovenia, and in the United States (Perz and Le Bancq, 2001; Ong et al., 2002; da Silva et al., 2003; Ryan et al., 2003a, 2005a; Learmonth et al., 2004; Leoni et al., 2006; Soba et al., 2006; Feltus et al., 2006; Blackburn et al., 2006). These findings suggest that oocysts of the Cryptosporidium cervine genotype excreted by sheep might be zoonotic.
IV.
Cryptosporidiosis in Goats
Cryptosporidium infection in goats was first described in a 2-week-old kid with diarrhea in Australia (Mason et al., 1981) and subsequently has been widely reported in many other areas (Kaminjolo et al., 1993; Abou-Eisha, 1994; Nagy, 1995; Morgan et al., 1998; Noordeen et al., 2000; Mahdi and Ali, 2002; ˇ ´ et al., 2006; Ozmen et Bomfim et al., 2005; Castro-Hermida et al., 2005; Delafosse et al., 2006; Misic al., 2006; Park et al., 2006). Cryptosporidiosis is considered an economically important disease, characterized by diarrhea with occasional death of kids. Several outbreaks have been reported. In Oman an outbreak resulted in the deaths of 238 goats, ranging from 2 days of age to over 1 year (Johnson et al., 1999). In Turkey and Brazil, outbreaks of cryptosporidiosis were responsible for diarrhea in 130 neonatal goat kids (Sevinç et al., 2005; Vieira et al., 1997). The histological appearance of infected intestinal
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tissue is similar to that described for other livestock, with widespread atrophy, blunting, and fusion of villi (Johnson et al., 2000). However, despite increasing recognition of the significance of cryptosporidiosis in goats, many features of the disease are poorly understood. Prevalence studies report infection rates of cryptosporidiosis in goats from 0 to 42.9% (e.g., Nagy, 1995; Muñoz et al., 1996; Majewska et al., 2000; Watanabe et al., 2005). Cryptosporidiosis has been ˇ ´ et al., detected in all ages of goats, although prevalence is higher in kids (Abou-Eisha, 1994; Misic 2006). A 12-month longitudinal study in Sri Lanka reported the highest prevalence (70.8%) at 2 months of age (Noordeen et al., 2001). The highest oocyst counts consistently were detected in the first few ˇ ´ et months of life, indicating that kids likely acquire infection as neonates (Noordeen et al., 2001; Misic al., 2006). Twelve kids experimentally infected with oocysts (from a calf with diarrhea) developed diarrhea; oocyst shedding started 4 days later and continued for 8 days (Koudela and Jii, 1997). A periparturient rise in oocyst excretion (Castro-Hermida et al., 2005) as well as long periods of oocyst shedding by asymptomatic animals (Noordeen et al., 2001) could serve as sources of infection for kids. Experimental infections of kids with oocysts from asymptomatic goats resembled natural infection in terms of oocyst excretion and clinical manifestation (Noordeen et al., 2002). Molecular data on Cryptosporidium species and genotypes in goats are very limited. Cryptosporidium parvum, C. hominis, and a goat genotype have been identified in natural infections in goats in Australia, Cyprus, the Czech Republic, Korea, Sri Lanka, Switzerland, and the United Kingdom (Morgan et al., ˇ et al., 2004; Ryan et al., 2005b; Park et 1998; Chalmers et al., 2002; Noordeen et al., 2002; Hajdusek al., 2006). Because most human infections are caused by C. hominis and C. parvum, both reported in goats, caprine cryptosporidiosis should be considered a potential zoonosis.
V.
Cryptosporidiosis in Pigs
The first reports of Cryptosporidium infection in pigs were published in 1977 (Bergeland, 1977; Kennedy et al., 1977). In South Dakota, histological examination of intestinal tissue from piglets with necrotic enteritis revealed a single piglet with Cryptosporidium stages developing on villous surfaces (Bergeland, 1977). In Kansas, tissue sections from three pigs, inoculated with colon contents from pigs with dysentery, also had stages of Cryptosporidium in the microvillus border of epithelial cells (Kennedy et al., 1977). Subsequently, cryptosporidiosis has been found in pigs, worldwide (Bergeland and Henry, 1982; Links, 1982; Sanford, 1987; Koudela et al., 1986; Tacal et al., 1987; Vitovec and Koudela, 1987; Xiao et al., 1994a; Kaminjolo et al., 1993; Guselle et al., 2003; Vitovec et al., 2006). The wide range of clinical features reported for cryptosporidiosis in pigs possibly reflects differences between gnotobiotic and conventionally raised piglets as well as differences among the Cryptosporidium isolates that caused infection (Table 18.11). Generally, prepatent periods range from 1 to 9 days and patent periods from 4 to 29 days. Clinical signs, including inappetance, depression, vomiting, and diarrhea, have been observed following experimental infection of piglets with oocysts derived from calves (Moon and Bemrick, 1981; Tzipori et al., 1981, 1982; Vitovec and Koudela, 1992; Enemark et al., 2003a). In five piglets with diarrhea Cryptosporidium was the only enteropathogen identified (Bergeland and Henry, 1982). However, cryptosporidiosis in pigs does not always result in clinical signs. One report on natural infections found significantly more diarrhea among infected pigs than in uninfected pigs (Hamnes et al., 2006), whereas other studies report no significant association between diarrhea and cryptosporidial infection (Sanford, 1987; Quílez et al., 1996a; Guselle et al., 2003; Maddox-Hyttel et al., 2006; Vitovec et al., 2006). In the absence of molecular analysis, is it not possible to determine which Cryptosporidium species or genotypes were responsible for the aforementioned infections. Isolates from diarrheic calves were most likely C. parvum, whereas natural infections could result from pigadapted species or genotypes. When species or genotype was known, infections from C. hominis, C. suis, or Cryptosporidium pig genotype II were less severe than those from C. parvum (Pereira et al., 2002; Enemark et al., 2003b; Guselle et al., 2003; Hamnes et al., 2006). A series of experimental coinfections with C. parvum and C. hominis demonstrated rapid displacement of C. hominis by C. parvum (Akiyoshi et al., 2003a). Clearly, identification of the species and genotypes is needed to establish the pathogenecity and host affinity of different Cryptosporidium isolates. Additionally, concomitant
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TABLE 18.11 Clinical Features of Cryptosporidium Infections in Pigs Source/Identity of Inoculum
Age
Type
Prepatent Period
Patent Period
Diarrhea
Infected calves
1d
Conventional 3–4 d
9–10 d
Yes, 9–11 d
Infected human (Immunodeficient) Infected calf Infected calf
Newborn Conventional 4–5 d
4–22 d
Several days
1d 1d
Gnotobiotic 1–4 d Conventional 3 d
NS 6d
Yes, w/vomiting Sporadic
Infected calf
1–2 d
Conventional 3–5 d
9d
Yes
Infected calf
1d
Gnotobiotic
3–5 d
16 d
Yes
C. hominis C. parvum C. meleagridis (isolated from a human) Infected calf
1d 1d Piglets
Gnotobiotic Gnotobiotic Gnotobiotic
8.6 d avg 5.6 d avg 4–5 d
16.6 d avg Mild 10.3 d avg Moderate 10–12 d Yes, but no dehydration
2d
C. suis
2d
Specific 3.3 d avg Pathogen Free Conventional 2–9 d
C. suis w/ concurrent 2 d natural rotavirus infection Natural infection (10 21–28 d samples typed as C. suis)
C. hominis
1d
Conventional 4–6 d
Conventional Mean time of initial oocyst excretion, post weaning 45 ± 16d Conventional 4–5 d
Reference Moon and Bemrick, 1981 Moon et al.,1982 Tzipori et al., 1982 Argenzio et al., 1990 Vitovec and Koudela, 1992 Vitovec and Koudela, 1992 Pereira et al., 2002 Pereira et al., 2002 Akiyoshi et al., 2003b
13.3 da avg Yes, 4.75 d avg Vomiting in 7 of 12 piglets 9–15 d Yes, 4 of 6 Vomiting in 2 of 6 piglets 13 db Yes, all—severe Vomiting in 2 of 6 piglets 29 d avg No
Guselle et al., 2003
4–10 d
Ebeid et al., 2003
Rarely
Enemark et al., 2003a Enemark et al., 2003b Enemark et al., 2003b
Note: d = day; NS = not specified. a b
Not all animals had stopped shedding at termination of observations. Data from 1 piglet only, remainder died or were euthanized 1 to 6 days after onset of symptoms because of severe illness.
infection with other pathogens such as Isospora suis or viruses can influence the severity of infections (Enemark et al., 2003b; Nuñez et al., 2003). For example, incidental coinfection with rotavirus apparently resulted in severe clinical signs and the deaths of five of six piglets experimentally infected with C. suis, whereas a rotavirus-free group had mild symptoms (Enemark et al., 2003b). Experimental infection with an avian species, C. meleagridis, consistently resulted in oocyst excretion and diarrhea, although mucosal changes were milder than those described for C. parvum (Akiyoshi et al., 2003b). Developing stages of Cryptosporidium have been differentially reported throughout the intestinal tract, with varying degrees of villous atrophy, villous fusion, cellular infiltration of the lamina propria and, infrequently, sloughing of epithelial cells (Kennedy et al., 1977; Moon and Bemrick, 1981; Links, 1982; Moon et al.,1982; Tzipori et al., 1982; Sanford, 1987; Koudela et al., 1989; Vitovec and Koudela, 1987, 1992; Argenzio et al., 1990; Pereira et al., 2002; Ebeid et al., 2003; Enemark et al., 2003a, 2003b; Nuñez et al., 2003; Vitovec et al., 2006). All species examined appear to cause similar types of histopathology, but lesions from C. parvum seemed most severe, reflected by the more significant clinical signs described for that species. Several histological studies have noted a “caudal shift.” Early in the infection, more stages were found in the upper intestine, whereas later, more stages were found in the lower regions (Tzipori et al., 1982; Vitovec and Koudela, 1992). Extraintestinal infections have been reported in the gall bladder of two naturally infected piglets (Fleta et al., 1995), and in the gall bladder, bile ducts, and
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pancreatic ducts of experimentally immunosuppressed piglets (Healey et al., 1997). Experimental inoculation produced infections in the trachea and conjunctiva of normal piglets (Heine et al., 1984a); the significance of infection at these extraintestinal sites is unknown. Detection and identification techniques are essentially identical to those described for cattle: microscopy, fecal antigen ELISA, and PCR, with or without prior application of a concentration technique. In comparing recovery and enumeration of C. parvum oocysts from feces of different animals, pig samples were the most problematic (Davies et al., 2003). Recovery of oocysts from pig feces was consistently lower than from feces of other animals. The authors suggested a possible problem with separation from immunomagnetic beads. Thus, it is not clear if pig feces present a problem with concentration techniques that do not employ immunomagnetic concentration methods. However, when analyzing fecal samples and reviewing data, it is important to recognize that cleaning and concentrating methods can be responsible, in part, for the variation seen in different prevalence studies. Serologic data also need to be interpreted with caution as they provide information on past infection status. In one prevalence study, 53% of adult pigs were serologically positive for Cryptosporidium, but none had detectable oocysts in their feces (Quílez et al., 1996c); in another study, 95% of 43 pig sera had antibodies to Cryptosporidium, considerably higher than the reported prevalences for fecal oocysts (Tzipori and Campbell, 1981). Definitive identification of species and genotypes is dependent on the use of molecular techniques, such as gene sequencing or PCR-RFLP. Given the ability of pigs to harbor multiple species/genotypes, some of which (C. parvum and C. suis) are infectious for humans, inclusion of molecular analysis in prevalence studies is necessary to understand Cryptosporidium populations in this host and to evaluate the risk of human infection. Point prevalence studies report infection rates from 0 to 88% of pigs tested, whereas a cumulative prevalence of 100% has been reported (see Table 18.12 for detailed references). The observed variation between studies, as discussed earlier, could be related to the variety of detection methods employed or the particular species/genotype to which the pigs are naturally exposed. Studies in which different age groups are sampled independently clearly demonstrate a relationship between Cryptosporidium prevalence and the age of the pigs. Cryptosporidium infections are most common in piglets greater than 1 month but generally less than 6 months of age (Fleta et al., 1995; Quílez et al., 1996a, 1996b, 1996c; ˇ ´ et al., 2003; Maddox-Hyttel et al., 2006; Vitovec et al., 2006); infections are Atwill et al., 1997; Misic detected less frequently in animals younger than 1 month or in adults (Sanford, 1983, 1987; Fleta et al., 1995; Quílez et al., 1996a, 1996b; Wieler et al., 2001; Hamnes et al., 2006; Maddox-Hyttel et al., 2006; Vitovec et al., 2006); and a number of studies have reported the absence of Cryptosporidium in these two age groups (Xiao et al., 1994a; Fleta et al., 1995; Quílez et al., 1996a, 1996b; Vitovec et al., 2006). Conventionally reared piglets less than 1 month of age have been reliably infected under experimental conditions (Moon and Bemrick, 1981; Argenzio et al., 1990; Vitovec and Koudela, 1992; Ebeid et al., 2003; Enemark et al., 2003a, 2003b), yet one longitudinal study reported that, on average, piglets began shedding oocysts at 45 days post weaning (Guselle et al., 2003). It is not clear why piglets under natural conditions appear to acquire infections when they are somewhat older. Natural infection of pigs with C. parvum, C. suis, and Cryptosporidium pig genotype II has been reported (Morgan et al., 1998, 1999; Guselle et al., 2003; Ryan et al., 2003b, 2004; Langkjær et al., 2007) (see Table 18.13), and as discussed earlier, experimental infections with C. hominis and C. meleagridis have been successful (Pereira et al., 2002; Akiyoshi et al., 2003a, 2003b; Enemark et al., 2003a, 2003b). The first evidence for a unique genotype of Cryptosporidium in pigs was reported in 1998 upon analysis of fecal samples from commercial pigs in Australia and Switzerland (Morgan et al., 1998), and feral pigs in California (Pereira et al., 1998). Further analysis of the “pig” genotype revealed that unlike the bovine isolates of Cryptosporidium, the pig genotype was not infectious for mice (Morgan et al., 1999), and upon additional molecular analysis, Cryptosporidium pig genotype I was elevated to species status as C. suis (Ryan et al., 2004). Subsequently, a second unique genotype was identified in pigs, Cryptosporidium pig genotype II (Ryan et al., 2003b; Hamnes et al., 2006; Langkjær et al., 2007; Xiao et al., 2006). Few prevalence studies have utilized molecular methods to analyze samples from pigs. In Canada, 33 piglets were monitored from weaning to market age (Guselle et al., 2003). All became infected with
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TABLE 18.12 Prevalence of Cryptosporidium in Pigs by Microscopy
Location Australia Australia Canada Canada Canada Canada Canada Czech Republic
Croatia Denmark
Germany Germany Ireland Italy Japan
Japan Korea Korea Norway Serbia
Spain
Spain Spain
Spain
Number of Animals/Number of Farms a
78 646/22 1453b 3491a 100/5 25–50c/1 33/1 8 farms 135 3368 835 38 production units/5 pigs each 49 farms 488 504 245 287f/24 1427a 56 pig slurry samples/33 farms 200 littersg 8 farms 232 252 108/slaughterhouse 500/NS 589/4 areas 684 littersg/100 # farms NS 50 40 38 45 42 45 Slaughterhouse 36 11 10 329i 27 farms 49 49 312 210 # Farms NS 14 16 30 30
Age
Prevalence (% positive)
NS <9w 5–15 d 1–30 w Varied Varied 21 d–6 m
0 6 0.5 5.3 11 0 100d
Sows Preweaned Postweaned NS
0 5.7 24.1 65.8e
<1m 1–4 m sows 1–42 d Varied Not applicable
6 71 4 1.4 0.1 50—single farm 40.6—other farms 0.5
Piglets
1–3 m 6m Adults 6–8 m Samples collected at slaughter 4–33 d
33.2 NS 0 19.6 62
1–30 d 1–3 m 3–6 m 6–9 m 9–12 m >12 m
32 62.5 44.7 31.1 23.8 15.5
1–2 m 2–6 m Adult < 2m
36h 9 0 3j
<1m 1–2 m 2–6 m Adult
0 59.2 34.3 0 + MZN 0 43.7 46.7 0
Reference O’Donoghue et al., 1987 Ryan et al., 2003b Sanford, 1983 Sanford, 1987 Olson et al., 1997b Heitman et al., 2002 Guselle et al., 2003 Vitovec et al., 2006
Bilic and Bilkei, 2006 Maddox-Hyttel et al., 2006
Wieler et al., 2001 Epe et al., 2004 Xiao et al., 2006a Canestri-Trotti et al., 1984a Izumiyama et al., 2001
Koyama et al., 2005 Rhee et al., 1991 Yu and Seo, 2004
8.3
Hamnes et al., 2006 ˇ ´ et al., 2003 Misic
Fleta et al., 1995
<1m 1–2 m 2–6 m Adult
Villacorta et al., 1991 Quílez et al., 1996a
+ IFA 0 87.5 56.7 0
Quílez et al., 1996b
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TABLE 18.12 (CONTINUED) Prevalence of Cryptosporidium in Pigs by Microscopy
Location Trinidad and Tobago Trinidad United States— California United States— California
United States— Ohio
Vietnam
Number of Animals/Number of Farms
Prevalence (% positive)
Age
Reference
275/NS
NS
19.6
Kaminjolo et al., 1993
52/3 200k
NS NS
1.9 5
Adesiyun et al., 2001 Tacal et al., 1987
10 feral populations 62 159 2 farms 44 litters 176 62 17/several
Atwill et al., 1997 8 ml 9m
11 3 Xiao et al., 1994a
7–25 d 5–8 w Sows 4–8 w
15.9% of littersm 8 0 23.5
Koudela et al., 1986
Note: d = day; w = week; m = month; NS = not specified; MZN = modified Ziehl-Nelson, acid-fast stain; IFA = immunofluorescent antibody stain. a b c d e f g h i j k l m
Animals/samples submitted to a diagnostic lab. Diarrheic animals submitted to a diagnostic lab. Authors state “25–50 samples were collected from all age groups present”; no additional explanation is provided. Longitudinal study, cumulative prevalence. Percent of units positive. Only diarrheic animals were sampled. Pooled fecal samples from a litter of piglets. Two piglets, 5.5%, had stages in the gall bladder. Various locations, farms, and slaughterhouses. All found in 1 litter of ten 50-day-old piglets. Stockyard animals. Determined by dentition. All on one farm.
Cryptosporidium, and isolates from ten were identified as C. suis (syn. pig genotype I). In a survey of 1237 pigs in Denmark, genotypic analysis was conducted on 170 and 13 microscopically positive samples from weaners and piglets, respectively (Langkjær et al., 2007). In weaners, 24% of the isolates were C. suis and 76% were Cryptosporidium pig genotype II, whereas in piglets, 71% of the isolates were C. suis and 29% were Cryptosporidium pig genotype II (Langkjær et al., 2007). Thus, it appears there could be age-related differences in prevalence. Pigs infected with C. suis produced more oocysts per gram of feces than those infected with pig genotype II; several pigs had mixed infections (Langkjær et al., 2007). In Norway, samples were collected from 100 swine herds, but only 9 of the microscopically positive samples were genotyped (Hamnes et al., 2006). Sequence analysis of the SSU-rRNA gene revealed six isolates were C. suis and three were pig genotype II. Analysis of pig slurry samples from 33 farms in Ireland revealed the presence of C. suis and Cryptosporidium pig genotype II, as well as C. muris (Xiao et al., 2006). It could not be determined if the C. muris originated in infected pigs or from farm rodents. Data thus far suggest there could be age-related differences in the prevalence of C. suis and the pig genotype II and that C. parvum is not widespread in pigs (Langkjær et al., 2007) (see Table 18.13). Sporadic cases of C. suis in humans have been reported (Xiao et al., 2002; Cama et al., 2003; Leoni et al., 2006), but the risk that pigs pose for human infections is unknown.
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TABLE 18.13 Cryptosporidium Species/Genotypes Reported from Pig with Molecular Data Cryptosporidium Species/Genotype (number of isolates)a
Country
Gene
Pig genotype I Pig genotype I (8) C. parvum (4) Pig genotype I (1) Pig genotype I (14) Pig genotype II (14) C. suis (5)
Australia Switzerland Australiab Switzerland Denmark Australia
SSU-rRNA SSU-rRNA
Morgan et al., 1998 Morgan et al., 1999
SSU-rRNA SSU-rRNA
Enemark et al., 2003b Ryan et al., 2003b
Australia Switzerland
Ryan et al., 2004
C. suis (10) C. suis (6) Pig genotype II (3) C. suis (50) Pig genotype II (133) C. suis (394) C. suis (16) Pig genotype II (11) C. muris (1)c
Canada Norway
SSU-rRNA Actin HSP-70 HSP-70 SSU-rRNA Actin SSU-rRNA HSP-70 SSU-rRNA SSU-rRNA
a b c
Demark Czech Republic Ireland
Reference
Guselle et al., 2003 Hamnes et al., 2006 Langkjær et al., 2007 Vítovec et al., 2006 Xiao et al., 2006
Pig genotype I was named as a new species, C. suis, by Ryan et al. (2004). One isolate identified as pig genotype I from Australia. Pig slurry samples; C. muris could have originated from either pigs or rodents.
VI. Cryptosporidiosis in Other Livestock Water buffalo (Bubalus sp.) are economically important animals raised for labor, meat, and milk in many parts of the world. Cryptosporidium oocysts were detected in feces of water buffalo in Italy, Brazil, Cuba, Egypt, and India (Canestri-Trotti and Quesada, 1983; Canestri-Trotti et al., 1984b; Galiero et al., 1994; Iskander et al., 1987; Rodríguez Diego et al., 1991; Dubey et al., 1992; Araujo et al., 1996). The prevalence of infection ranged from 6 to 39%. One study found a higher incidence of infection in diarrheic buffalo; another found no conclusive relationship (Dubey et al., 1992; Galiero et al., 1994). Of 616 and 525 buffalo examined in Cuba and Egypt, respectively, oocysts were found only in calves (Abou-Eisha, 1994; Rodríguez Diego et al., 1991). Llamas (Lama glama) and alpacas (Lama pacos) are usually raised as work animals and for fiber production, respectively. Cryptosporidiosis has been occasionally detected in both species. Analysis of diarrheic feces collected from preweaned llama and alpaca crias in Oregon, revealed that 9% were Cryptosporidium positive (Cebra et al., 2003). Of 354 llamas examined in California, no oocysts were detected (Rulofson et al., 2001). There is a single report of C. parvum, confirmed by molecular analysis, in an alpaca from Peru (Morgan et al., 1998). Camels (Camelus bactrianus and C. dromedarius) are widely used for food, fiber, and as work animals across parts of Asia and Africa. Most reports of Cryptosporidium in camels have been from zoo-housed animals (Fayer et al., 1991; Abou-Eisha, 1994; Gómez et al., 2000; Gracenea et al., 2000). Cryptosporidium was detected 1 of 4 camel calves in Egypt (Abou-Eisha, 1994) but in none of 23 camels from Iraq (Mahdi and Ali, 2002). Oocysts recovered from a zoo-housed camel (Fayer et al., 1991) were infectious for mice (Anderson, 1991) and were determined to be C. muris (Xiao et al., 1999; Morgan et al., 2000), whereas oocysts from camels in the Czech Republic were typed as C. andersoni. The American Bison (Bison bison) sometimes occupies grazing lands on or adjacent to those occupied by beef cattle, and some bison are raised for meat in the United States and Canada. Cryptosporidium has been detected in both the American and European bison (B. bonasus) at the Lisbon and Barcelona
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Zoos (Gómez et al., 2000; Alves et al., 2005). Surprisingly, the isolate from the American Bison at the Lisbon Zoo was Cryptosporidium mouse genotype (Alves et al., 2005).
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Tzipori, S., Rand, W., Griffiths, J., Widmer, G. and Crabb, J. 1994. Evaluation of an animal model system for cryptosporidiosis: therapeutic efficacy of paromomycin and hyperimmune bovine colostrumimmunoglobulin. Clin. Diagn. Lab. Immunol. 1: 450–463. Uehlinger, F.D., Barkema, H.W., Dixon, B.R., Coklin, T. and O’Handley, R.M. 2006. Giardia duodenalis and Cryptosporidium in a veterinary college bovine teaching herd. Vet. Parasitol. 142: 231–237. Uga, S., Matsuo, J., Kono, E., Kimura, K., Inoue, M., Rai, S.K. and Ono, K. 2000. Prevalence of Cryptosporidium parvum infection and pattern of oocysts shedding in calves in Japan. Vet. Parasitol. 94: 27–32. Ungar, B.L., Kao, T.C., Burris, J.A. and Finkelman, F.D. 1991. Cryptosporidium infection in an adult mouse model. Independent roles for IFN-gamma and CD4+ T lymphocytes in protective immunity. J. Immunol. 147: 1014–1022. Vieira, L.S., Silva, M.B., Tolentino, A.C., Lima, J.D. and Silva, A.C. 1997. Outbreak of cryptosporidiosis in dairy goats in Brazil. Vet. Rec. 140: 427–428. Villacorta, I., Ares-Mazas, E. and Lorenzo, M.J. 1991. Cryptosporidium parvum in cattle, sheep and pigs in Galicia (N.W. Spain). Vet. Parasitol. 38: 249–252. Vitovec, J. and Koudela, B. 1987. Natural cryptosporidiosis in a piglet suffering from pseudomembranous typhlitis. Folia Parasitol. 34: 378. Vitovec, J. and Koudela, B. 1992. Pathogenesis of intestinal cryptosporidiosis in conventional and gnotobiotic piglets. Vet. Parasitol. 43: 25–36. Vitovec, J., Hamadejova, K., Landova, L., Kvac, M., Kvetonova, D. and Sak, B. 2006. Prevalence and pathogenicity of Cryptosporidium suis in pre- and post-weaned pigs. J. Vet. Med. B Infect. Dis. Vet. Pub. Health. 53: 239–243. Wade, S.E. Mohammed, H.O. and Schaaf, S.L. 2000. Prevalence of Giardia sp. and Cryptosporidium parvum and Cryptrosporidium muris (C. andersoni) in 109 dairy herds in five counties in southeastern New York. Vet. Parasitol. 93: 1–11. Watanabe, Y., Yang, C.H. and Ooi, H.K. 2005. Cryptosporidium infection in livestock and first identification of Cryptosporidium parvum genotype in cattle feces in Taiwan. Parasitol. Res. 97: 238–241. Wee, S.H., Joo, H.D. and Kang, Y.B. 1996. Evaluation for detection of Cryptosporidium oocysts in diarrheal feces of calves. Korean J. Parasitol. 34: 121–126. Wieler, L.H., Ilieff, A., Herbst, W., Bauer, C., Vieler, E., Bauerfeind, R., Failing, K., Klös, H., Baljer, G. and Zahner, H. 2001. Prevalence of enteropathogens in suckling and weaned piglets with diarrhoea in Southern Germany. J. Vet. Med. B 48: 151–159. Wu, Z., Nagano, I., Boonmars, T., Nakada, T. and Takahashi, Y. 2003. Intraspecies polymorphism of Cryptosporidium parvum revealed by PCR-restriction fragment length polymorphism (RFLP) and RFLP-singlestrand conformational polymorphism analyses. Appl. Environ. Microbiol. 69: 4720–4726. Yu, J.R., Lee, J.K., Seo, M., Kim, S.I., Sohn, W.M., Huh, S., Choi, H.Y. and Kim, T.S. 2004. Prevalence of cryptosporidiosis among the villagers and domestic animals in several rural areas of Korea. Korean J. Parasitol. 42: 1–6. Yu, J.R. and Seo, M. 2004. Infection staus of pigs with Cryptosporidium parvum. Korean J. Parasitol. 42: 45–47. Xiao, L. and Herd, R.P. 1994. Infection pattern of Cryptosporidium and Giardia in calves. Vet. Parasitol. 55: 257–262. Xiao, L. and Ryan, U.M. 2004. Cryptosporidiosis: an update in molecular epidemiology. Curr. Opin. Infect. Dis. 17: 483–490. Xiao, L., Herd, R. and Rings, D. 1993. Diagnosis of Cryptosporidium on a sheep farm with neonatal diarrhea by immunofluorescence assays. Vet. Parasitol. 47: 17–23. Xiao, L., Herd, R.P. and Bowman, G.L. 1994a. Prevalence of Cryptosporidium and Giardia infections on two Ohio pig farms with different management systems. Vet. Parasitol. 52: 331–336. Xiao, L., Herd, R.P. and McClure, K.E. 1994b. Periparturient rise in the excretion of Giardia cysts and Cryptosporidium parvum oocysts as a source of infection for lambs. J. Parasitol. 80: 55–59. Xiao, L., Escalante, L., Yang, C., Sulaiman, I., Escalante, A.A., Montali, R.J., Fayer, R., Lal, A.A. 1999. Phylogenetic analysis of Cryptosporidium parasites based on the small subunit rRNA gene locus. Appl. Enviroon. Microbiol. 65: 1578–1583. Xiao, L., Fayer, R., Ryan, U., Upton, S.J., 2004. Cryptosporidium taxonomy: recent advances and implications for public health. Clin. Microbiol. Rev. 17: 72–97.
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Xiao, L., Moore J.E., Ukoh, U., Gatei, W., Lowery, C.J., Murphy, T.M., Dooley, J.S., Millar, B.C., Rooney, P.J., and Rao, J.R. 2006. Prevalence and identity of Cryptosporidium in pig slurry. Appl. Environ. Microbiol. 72: 4461–4463. Xiao, L., Zhou, L., Santín, M., Yang, W. and Fayer, R. 2007. Distribution of Cryptosporidium parvum subtypes in calves in eastern United States. Parasitol. Res. 100: 701–706 Xiao, L., Sulaiman, I.M., Ryan, U.M., Zhou, L., Atwill, E.R., Tischler, M.L., Zhang, X., Fayer, R. and 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. Zarlenga, D.S. and Trout, J.M. 2004. Concentrating, purifying and detecting waterborne parasites. Vet. Parasitol. 126: 195–217.
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19 Animal Models
Saul Tzipori and Giovanni Widmer
CONTENTS I. II.
Introduction .................................................................................................................................. 485 Animal Models for Parasite Propagation and Oocyst Production .............................................. 486 A. Introduction ..................................................................................................................... 486 B. Generation of C. parvum Oocysts in Calves.................................................................. 486 C. Generation of C. hominis Oocysts from Piglets............................................................. 487 III. Traditional and New Animal Models for Research..................................................................... 487 A. Introduction ..................................................................................................................... 487 B. Models to Evaluate Agents for Treatment or Prevention............................................... 488 1. Overview .......................................................................................................... 488 2. The Neonatal Mouse Model ............................................................................ 488 3. The Immunosuppressed Rodent Models ......................................................... 489 4. The Gnotobiotic Piglet Diarrhea Model.......................................................... 489 5. The SIV-Infected Rhesus Monkey Model....................................................... 490 6. The Gerbil Model ............................................................................................ 490 Cryptosporidium Genetics .............................................................................................. 491 C. D. Testing of Water Treatments ........................................................................................... 492 E. Animal Models versus Cell Culture ............................................................................... 492 IV. Animal Models and Cryptosporidium Host Range ..................................................................... 493 References........................................................................................................................... 493
I.
Introduction
Since the discovery of the Cryptosporidium genus some 100 years ago, the field went through several metamorphoses. Based on early cross-species transmission experiments, investigators had concluded that, similar to other members of the Apicomplexa such as malaria and coccidia, Cryptosporidium was a multispecies genus (Levine, 1984). Thus, isolates were named after the species of animal in which they were observed, and by 1980 the list consisted of some 23 species, which included C. bovis, C. hominis, C. ovis, and so on (Tzipori, 1983). In 1980, in a series of experiments conducted at the Moredun Institute in Scotland, it was shown that the predominant Cryptosporidium species that perpetuated among all the domestic animals and humans lacked host specificity (Tzipori et al., 1980). Over the following two-and-a-half decades, using sophisticated genetic tools, differentiation of species of the genus Cryptosporidium made gradual progress. However, the discovery in 1980 that Cryptosporidium isolated from cattle was capable of infecting a wide range of domestic mammals, including rodents (Tzipori et al., 1980), opened up the field to the use of rodents in early animal experiments. These experiments included testing of antimicrobial agents (Tzipori et al., 1982b), disinfectants (Campbell et al., 1982), seroprevalence of the infection in key domestic and human species (Tzipori and Campbell, 1981), and
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many cross-transmission experiments (Tzipori, 1983). These early observations were subsequently confirmed by others (Reese et al., 1982). The use of rodents has enormously advanced our knowledge in this field, and until the end of the 20th century, the bulk of animal experiments used C. parvum propagated largely in calves as the source of parasite. This situation has changed, however, with the recognition that C. parvum in fact consists of two genetically and phenotypically distinct species (Carraway et al., 1996; Morgan-Ryan et al., 2002; Peng et al., 1997). Whereas C. parvum continues to be by far the predominant species infecting many mammals, C. hominis in nature infects humans almost exclusively. Although C. hominis does infect other mammalian species under laboratory conditions, including calves, piglets, and gerbils (see following text), the species’ inability to infect other more commonly used rodents is a major setback, and has considerably limited the extent of research that can be performed with C. hominis. Animal models for cryptosporidiosis are required for the generation of oocysts, the exogenous infectious forms used for laboratory investigations, and models in which to perform such studies. Curiously, despite the availability of several sensitive models such as the INF- knockout (GKO) mouse, most investigators use either the neonatal, or the immunosuppressed mouse.
II. A.
Animal Models for Parasite Propagation and Oocyst Production Introduction
The inability to continuously propagate C. parvum in vitro remains the most important obstacle to performing the kind of studies that are possible with so many other pathogens of major interest. This limitation is reflected in the lack of access to stages of the life cycle other than sporozoites and the oocysts, and the need to use animals to generate oocysts for research. Furthermore, oocysts generated in animals have a limited lifespan. For instance, infectivity markedly diminishes within 6 to 8 weeks for in vitro and in vivo infectivity studies. Older oocysts may, however, still be useful for animal propagation, and as a source of antigen for serology, biochemistry, and for extracting nucleic acid for genetic work. As there are presently no methods that allow indefinite storage of infectious material, isolates have to be continuously passaged through animals, usually calves or mice for C. parvum, and piglets or gerbils for C. hominis (see below). Oocysts and sporozoites are the only stages of the parasite life cycle that can be produced in large quantities in experimentally infected calves. Oocysts are readily purified and concentrated from calf feces. Oocysts can also be obtained from small ruminants experimentally infected immediately after birth (Hill et al., 1990; Tilley and Upton, 1991). Some investigators use rodents to generate small quantities of oocysts (Hill et al., 1991; Suresh and Rehg, 1996). Another major source of material used to be available from chronically infected human patients with AIDS (Giacometti et al., 1996; Morgan et al., 2000; Widmer et al., 1998). This source, however, is no longer available in the developed world since the introduction of retroviral therapy against HIV. Human patients tend to provide small and variable quantities of material, which makes referenced, comparative studies on genetic variation, virulence, and host susceptibility impossible.
B.
Generation of C. parvum Oocysts in Calves
Calves are infected immediately after birth, preferably before feeding on colostrum. They should be housed in complete isolation to protect them from exposure to other pathogens, including C. parvum from other sources, and to protect the individuals handling the infected animals. Infected calves constitute the greatest risk to humans because of the large number of oocysts generated by them. There are several published reports of animal handlers (Blagburn and Current, 1983) and veterinary students (Anderson and Bulgin, 1981) who became infected through contact with infected calves. Infected calves are kept in specially designed stanchions once diarrhea begins to facilitate the collection of fecal material. During the course of oocyst shedding, calves are fed an enriched calf electrolyte formula that facilitates the purification of oocysts by decreasing fecal fat and roughage content.
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There are several purification, concentration, and sterilization procedures, some of which were reviewed (Current, 1990). The method we have adopted in our laboratory combines several of them. Briefly, homogenized and filtered calf feces are mixed with saturated NaCl solution and centrifuged at 1000 × g for 15 min at 4˚C. The supernatant containing the oocysts is removed and diluted 1:3 in distilled water, and centrifuged at 4000 × g for 15 min. The pellet is resuspended and washed in 0.1% thiosulfate solution before pelleting and resuspending in Alsever’s solution for Percoll gradient centrifugation (Arrowood and Sterling, 1987; Waldman et al., 1986). After counting in a hemocytometer, oocysts are stored at 4˚C in antibiotic solution and are used within 6 to 8 weeks for infectivity studies. Oocysts are surface-sterilized in 1.75% sodium hypochlorite for 7 min before using them for in vitro or in vivo studies. Using these and similar methods, it is possible to obtain up to 1010 oocysts from a single calf. Sedimentation of fecal samples on step gradients of Histodenz (Sigma) has also been found effective for purifying oocysts from small volumes of fecal concentrates or feces (Widmer et al., 1998).
C.
Generation of C. hominis Oocysts from Piglets
Although oocysts of C. hominis can also be produced in gerbils (Baishanbo et al., 2005) and in calves (Akiyoshi et al., 2002), the risk of contamination of C. hominis isolates with C. parvum limits the use of such animals, particularly of calves, for long-term maintenance of pure C. hominis isolates. Hence, to maintain homogeneity and purity of C. hominis, it was found necessary to propagate C. hominis in germ-free animals. The human isolate TU502, which was used for sequencing the C. hominis genome (Xu et al., 2004), has been maintained in gnotobiotic piglets for over 5 years (Akiyoshi et al., 2002). In addition, it has been characterized with respect to its infectivity to human adult volunteers (Chappell et al., 2006), and is probably the only isolate of this species that can be considered a reference isolate. Unlike the currently used C. parvum isolates, which are propagated in conventional, farm-derived calves, such as IOWA (Abrahamsen et al., 2004), MD (Okhuysen et al., 2002), and others, which at some point may have become contaminated with farm isolates, C. hominis isolates of human origin such as TU502 and NEMC1 (Widmer et al., 2000) remained free of C. parvum and showed no sign of genotypic changes following serial propagation in germ-free piglets.
III. Traditional and New Animal Models for Research A.
Introduction
A review of the literature shows that currently most animal models are used for the evaluation of compounds for inhibitory activities against cryptosporidiosis. Although several sensitive rodent models have been developed, standardized, and optimized over the last two decades (see following text), the majority of investigators, presumably for reasons of availability and costs, continue to use either the neonatal mouse or the immunosuppressed adult mouse, which have major limitations. Although cryptosporidiosis occurs in most vertebrates, human cryptosporidiosis is quite distinct from that seen in other mammals. The infection in humans induces serious symptoms of gastrointestinal illness lasting several days in all age groups. Whereas infections cause acute watery diarrhea only in neonatal ruminants and piglets, no symptoms are observed in most other infected animals of any age. Because of these differences, different animal models are often required for different types of scientific studies: e.g., newborn calves are useful for studying pathogenesis and pathophysiology of the gut; mice to delineate specific aspects of the immune response; macaques are used to study the chronic disease in the immunodeficient host; piglets have been recently used to study C. hominis, etc. Because of the significance of cryptosporidiosis in other mammals, namely, calves and small ruminants, studies have been conducted since 1980 to investigate the infections and diseases that are unique to economically important domestic animals (O’Donoghue, 1995; Tzipori, 1983). The pathophysiologic impact of cryptosporidiosis on the entire host (Argenzio et al., 1990; Argenzio et al., 1993), as well as at specific gastrointestinal sites (Moore et al., 1995; Tzipori and Griffiths, 1998), was investigated using piglets or calves. For immunological and immunopathological studies, the mouse
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(Chen et al., 1993a, 1993b; Riggs, 2002; Theodos et al., 1997; Ungar et al., 1990, 1991), the SIV-infected monkey (Singh and Tzipori, manuscript in preparation), and other neonates, including lambs (Hill et al., 1990) and calves (Fayer et al., 1989; Peeters et al., 1992) were used. These studies provided valuable information about cryptosporidiosis in general, as well as information related to specific animal species (Current and Garcia, 1991; O’Donoghue, 1995; Tzipori, 1988, 1998).
B. 1.
Models to Evaluate Agents for Treatment or Prevention Overview
Although the pressure to develop effective drugs against cryptosporidiosis to combat persistent infections in individuals with HIV/AIDS (the main target population during the 1980s and 1990s) has subsided with the introduction of antiretroviral therapy, the need for therapy against the disease in humans and calves is as urgent as it has ever been. Effective therapy is urgently needed against acute infections in humans, in whom cryptosporidiosis tends to be more severe than most other forms of diarrhea, perhaps with the exception of shigellosis, and against the risk of persistent infections in immunosuppressed, immunodeficient, the elderly, and young children. Because the search for an effective therapy against persistent cryptosporidiosis remains the major objective of some drug development programs, the use of an immunodeficient host is highly desirable. The ideal immunocompromised animal model should (1) allow a rapid and consistent establishment of persistent infection in adult animals; (2) require a lowto-moderate infectious dose; and (3) generate a profound watery diarrhea, dehydration, malabsorption, weight loss, and emaciation. Moreover, it is critical that the infection colonize a major portion of the gastrointestinal tract and have the ability to spread to the hepatobiliary system, resulting in morphological changes similar to those seen in humans. There are several reported rodent models that are being used to screen drugs for their efficacy against C. parvum infection, none of which, however, meet all of the aforementioned criteria. These include (1) the normal neonatal mouse (Fayer and Ellis, 1993) or the neonatal scid mouse (Rohlman et al., 1993; Tzipori et al., 1994), (2) the dexamethasone (DEX) immunosuppressed adult mouse model (Yang and Healey, 1994), (3) the immunosuppressed rat model (Brasseur et al., 1994; Rehg et al., 1987), (4) the adult scid mouse model (Mead et al., 1991), and (5) the anti-IFNγ conditioned weaned scid model (Tzipori et al., 1995). Each model offers its own advantages and limitations. A disadvantage to the immunosuppressed adult mouse and rat models is that these animals will resolve their C. parvum infection unless immunosuppression is maintained. This often creates potential toxicity problems because administration of an immunosuppressive drug could possibly result in untoward interactions between the experimental and the immunosuppressive drugs. An added disadvantage of rats is that larger amounts of drugs are required as compared to a murine model, owing to differences in size. Although the adult scid mouse offers an immunocompromised model that can be easily randomized and manipulated, it takes a minimum time of 2 to 3 weeks before these mice start to shed detectable levels of oocysts (Mead et al., 1991; Tzipori et al., 1995). In addition, although the hepatobiliary system does become infected in adult scid mice, it generally takes 12 to 13 weeks before extraintestinal infections are consistently observed in all mice. This lengthens the time that mice need to be maintained for a drug trial, thereby increasing the expense.
2.
The Neonatal Mouse Model
The infant mouse, aged 4 to 6 days, is highly susceptible to infection and can be challenged with fewer than 100 oocysts (Finch et al., 1993; Korich et al., 2000). They remain clinically healthy and continue to shed oocysts for 10 to 14 days, after which they fully recover and stop shedding oocysts. Neonates invariably are used for assessing preventive treatments, rather than therapy, because of the short window of infection. Litters of ICR mouse pups are randomized into the desired groups on day 5, one day prior to challenge. Normally, treatment commences when mice are challenged. The following is a brief outline of the model used in our laboratory, which differs somewhat from those described by others (Blagburn et al., 1991; Fayer et al., 1990). The course of treatment is 5 to 6 days, involving two oral dosings of 30 to 50 µL volumes per dose by gavage. Mice are euthanized 18 h after the end of treatment, and the entire gastrointestinal tract is removed, weighed, and homogenized 1:40 (weight/volume) in PBS. The
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number of oocysts present in a given volume is counted in a hemocytometer chamber. The enumeration is repeated five times per mouse and averaged. The mean (±SD) per treatment group is calculated and expressed as percent inhibition as compared with the placebo. Paromomycin is normally used as the positive control drug. Among the limitations of using the infant mouse are the small volume that can be delivered accurately into the esophagus without inflicting injury, the short treatment period, variation in weight among litters, the potential for maternal rejection after randomization and human handling, and the fact that this model is limited to testing drugs prophylactic.
3.
The Immunosuppressed Rodent Models
Several methods to induce immunosuppression in rodents have been described, of which the administration of DEX in drinking water to either adult C57BL/6 mice (Yang et al., 1996) or to adult female rats (Rehg et al., 1988) is the most convenient. Immunosuppressed C57BL/6 mice given 10 to 100 μg/mL DEX in drinking water were challenged with 106 oocysts/mouse on day 14 post immunosuppression. With a dose of 10 μg/mL, mice began to shed oocysts 3 days post infection and continued for 45 days before succumbing to drug toxicity, a sufficient window for drug testing. It was not clear whether hepatobiliary tract infection occurred. The rat model only requires 0.25 μg/mL of DEX for 10 days to become immunosuppressed (Rehg et al., 1988). Rats were challenged with 106 oocysts, and by day 21 oocysts were detected in the feces of 75% of infected rats, and bile duct infection was evident in 68% of the animals by day 30. A drawback of all methods of immunosuppression, however, is the associated mortality level and other health complications.
4.
The Gnotobiotic Piglet Diarrhea Model
This model has a dual purpose. It is used extensively in the laboratory to propagate C. hominis to generate oocysts, and to characterize new C. hominis isolates (Akiyoshi et al., 2002; Pereira et al., 2002), as well as to evaluate therapeutic agents that have shown considerable activities in the cell culture or in a rodent model (Theodos et al., 1998; Tzipori et al., 1994). The obvious advantage of this model is the development of acute watery diarrhea and other gastrointestinal symptoms, which rodent models lack (Tzipori, 1983). The manifestation of diarrhea, a key symptom in human patients, is essential for the evaluation of oral therapeutic agents. Severe diarrhea affects drug transit time through the gastrointestinal tract and can markedly reduce their therapeutic action (Tzipori et al., 1994). Colostrum-deprived newborn piglets are highly susceptible to cryptosporidiosis, which leads to watery diarrhea, dehydration, malabsorption, wasting, and often sudden death within 2 to 5 days of onset of symptoms (Tzipori et al., 1982a, 1994; Vitovec and Koudela, 1992). In contrast, piglets raised under conventional conditions rarely experience diarrhea due to this infection, although cryptosporidiosis in swine is extremely prevalent, as judged by serological (Quilez et al., 1996) and microbiological surveys (Guselle et al., 2003; Sanford, 1987). Swine, as do cattle (Lorenzo et al., 1993; Scott et al., 1995), continue to intermittently excrete oocysts in their feces for long periods of time. Different Cryptosporidium spp. are involved in natural infections in these animals. Furthermore, it is not clear whether these animals regularly become reinfected or if they maintain a low-grade but persistent infection. Even colostrum-deprived piglets become rapidly resistant to C. parvum, and infection becomes mostly asymptomatic 7 to 10 days after birth. It is thought that because of the widespread and high prevalence of Cryptospridium (Quilez et al., 1996), good maternal protection is afforded to piglets during those first few days after birth. Studies have shown that the majority of pigs become subclinically infected shortly after weaning when aged 4 to 5 weeks (Quilez et al., 1996; Vanopdenbosch and Wellemans, 1985). Because colostrum-deprived newborn piglets are highly susceptible to infection with many other pathogens, and to avoid cross-infection among experimental groups, cesarean-derived gnotobiotic piglets maintained in complete isolation are used for drug evaluation. They are divided into groups and challenged 24 h after birth. Drug treatment normally begins on day 3 with the onset of watery diarrhea and oocyst excretion in feces. Treatment is either mixed in the milk diet or orally administered over 10 to 11 days. Parameters used for assessment of therapy include gastrointestinal symptoms, including severity of diarrhea and extent of illness, weight gain/loss, oocyst shedding, and degree of mucosal damage and infection at the end of treatment (Tzipori et al., 1994).
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The piglet diarrhea model helps identify realistic interactions between diarrhea, effective therapy, and drug toxicity. It highlights the impact of the severity of diarrhea on the outcome of treatment: the more severe the diarrheal illness, which accelerates gut transit by the drug, the poorer the clinical response to treatment with paromomycin is (Tzipori et al., 1994). These studies also showed the detrimental effect of stomach acidity on the efficacy of hyperimmune bovine colostrum and paromomycin. Finally, in the pig, individual variation in response to treatment can, as in human patients, be assessed in the context of the extent of infection and severity of illness. The limitations of this model include (1) variation in litter size (8 to 12 piglets), and variation in piglet size (0.8 to 2.0 kg); (2) the infection and illness, although they can be severe, are self-limiting, as animals recover with 14 days; (3) the overall susceptibility of piglets can vary greatly, from being highly susceptible, resulting in mortality, to being mildly infected and recovering rapidly, even when using the same oocysts as inoculum; and (4) the model is labor intensive and expensive as compared to rodents. It remains, however, the only standard diarrhea model available that can be used under controlled laboratory conditions.
5.
The SIV-Infected Rhesus Monkey Model
The ultimate model, however, that mimics many of the disease aspects observed in humans with AIDS is a primate with AIDS (Baskerville et al., 1991; Blanchard et al., 1987; Kaup et al., 1994). Immunocompetent primates develop a self-limiting acute diarrhea that can be life threatening in the young (Riggs, 1990, 2002). Progressive, fatal illness was described in macaques infected with the simian immunodeficiency virus (SIV) and cryptosporidiosis involving the gastrointestinal tract, complicated by hepatobiliary and respiratory tract infections (King, 1986; Osborn et al., 1984; Riggs, 1990). However, the expense, limited availability, and many other issues, including ethical considerations, limit extensive use of this model. There are, however, pertinent aspects of the disease and intervention that can only truly be investigated in such animals. Among them is the delineation of the various immunological defects in the gastrointestinal mucosa associated with progression of SIV infection that contribute to persistence of infection and to mucosal damage during cryptosporidiosis.
6.
The Gerbil Model
The discovery that immunosuppressed gerbils are susceptible to C. hominis is one of the most significant recent innovations in animal model research relevant to Cryptosporidium (Baishanbo et al., 2005). This finding was unexpected, as until recently C. hominis was not thought to be infectious for rodents (MorganRyan et al., 2002). This discovery removes an important obstacle to those working on this species. Although in most parts of the world C. parvum is not as prevalent as C. hominis in human infections (Thompson et al., 2005), C. parvum is commonly used as a surrogate owing to the relative ease with which the parasite can be obtained and grown in rodents. Until the successful infection of gerbils with C. hominis was reported, research on this species was dependent on oocysts isolated from human patients or on highly specialized animal models, such as the gnotobiotic pig (Tzipori et al., 1994). Baishanbo et al. (2005) successfully used 4- to 5-week-old gerbils (Meriones unguiculatus) that were immunosuppressed with 0.8 mg DEX (20 to 50 μg/kg) injected every second day starting 10 days before infection until day 21 post infection. They found that the oral administration of 100 oocysts was sufficient to generate sustained infections in DEX-treated gerbils. Although the gerbil is not as commonly used as other rodents in infectious disease research, it has emerged as a surprisingly versatile model. Its applications include research on trematodes (Chisty et al., 2002), cestodes (Sato et al., 2000), nematodes (Ostlind et al., 1990), protozoa (Campbell et al., 1999; Franco et al., 1991; Schaefer et al., 1991), and as a model for Helicobacter pylori-induced gastric ulcer (Fujioka et al., 2000). Although the gerbil model will greatly facilitate research on Cryptosporidium species, it suffers from two limitations: a lack of immunological reagents, and the absence of symptoms. Compared to the more commonly used rats and mice, few antibodies are available for exploring the immune response in this species. We have repeated the observations of Baishanbo et al. (2005) using three C. hominis isolates, including the TU502 isolate, which was used for sequencing the genome of C. hominis (Xu et al., 2004), and have extended them to two additional species, C. muris and C. meleagridis (Widmer, unpublished). Gerbils were injected intramuscularly with 1 mg of DEX (Sigma cat. no. D1756) every second day on alternate
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1 mg
160 µg/ml
MD
E
mouse 6×104
-6
-4
-2
0
TU71
2
4
6
8
10
12
14
16
E
gerbil 4×103
TU728 pig 2.5×104
TU502 gerbil 1.7×105
E
E
FIGURE 19.1 Experimental infections of gerbils with C. hominis and C. parvum. Four groups of four 4-week-old gerbils (Charles River Laboratories, Wilmington, MA) were immunosuppressed with dexamethasone starting on day 6 prior to infection (day 6 postinfection) and infected on day 0 (hatched squares) with C. parvum isolate MD (Okhuysen et al., 2002), or C. hominis isolate TU71, TU728 (Tanriverdi and Widmer, 2006), or TU502 (Akiyoshi et al., 2002; Xu et al., 2004) as indicated at extreme left. The host origin and number of oocysts orally administered are indicated below each isolate code. Each day is indicated by a square and labeled with respect of the day of infection (day 0) as shown for the MD isolate. The animals were immunosuppressed with four intramuscular injections of 1 mg dexamethasone as indicated by the syringes. Starting on day 3 postinfection, dexamethasone was added to the drinking water at a concentration of 160 μg/mL as represented by a bottle and horizontal arrow. Feces were monitored by acid-fast staining for the presence of oocysts starting on day 4 postinfection. Presence of oocysts in the feces is indicated with color (green, C. parvum; red, C. hominis). E indicates euthanasia.
hind legs starting on day 6 before infection. Gerbils were infected with various doses of C. hominis or C. parvum oocysts administered orally as shown in Figure 19.1. Starting on day 3 post infection, dexamethasone 21-phosphate disodium salt (Sigma cat. no. D-1159) was added to the drinking water at a concentration of 160 μg/mL. This solution was replaced every second day. Oocyst shedding typically started on day 7 post infection for C. hominis, and a few days earlier, usually on day 4, for C. parvum. Oocysts excreted by C. hominis-, C. parvum-, and C. meleagridis-infected gerbils were genotyped to exclude contaminations with different species. Immunosuppression protocols were not further optimized, and it is possible that the amount of injected DEX and the concentration of this compound in the water could be reduced to limit morbidity, and occasional mortality, without compromising susceptibility to C. hominis. The ID50 value for C. hominis in this model has not been determined, but results obtained by Baishanbo et al. (2005) suggest that it is less than 100 oocysts, which would be similar to that of C. parvum in mice (Finch et al., 1993; Korich et al., 2000).
C.
Cryptosporidium Genetics
The mouse model has enabled the application of genetic methods to the study of C. parvum. Although the differentiation of gametes, fertilization, and meiosis have been documented with electron microscopy (Goebel and Braendler, 1982; Tzipori, 1988), experimental evidence for the existence of a sexual phase in the life cycle of C. parvum is more recent (Feng et al., 2002). Experiments in which two genetically distinct isolates were crossed in GKO mice confirmed what was assumed on the basis of the life cycle in other apicomplexans, and in particular from Plasmodium species, namely, that sexual reproduction and genetic recombination is a part of the Cryptosporidium life cycle. The inability to positively select for C. parvum recombinants and exclude parental genotypes makes genetic studies labor intensive. Progeny oocysts obtained from mice coinfected with two isolates need to be propagated individually until clonal lines carrying alleles from both parental isolates are obtained. GKO mice were used for this task because they can be infected with a single oocyst (Tanriverdi et al., 2007). The observation that recombinant progeny lines can be more virulent than the parental isolates from which they were derived
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(Feng et al., 2002) indicates that virulence in C. parvum is encoded by multiple genetic loci. One of the goals of these studies is to identify such genes. We anticipate that the gerbil model will enable similar studies in C. hominis.
D.
Testing of Water Treatments
Although now mostly superseded by cell culture, the neonatal mouse model was used extensively to assess the sensitivity of C. parvum oocysts to water disinfectants (Korich et al., 1990; Peeters et al., 1989). Initially, the proportion of mice that became infected following oral inoculation with a dose of treated or control oocysts was used to quantify oocyst sensitivity to chlorine, chloramine, monochloramine, and ozone. Later, a dose-response design (Finch et al., 1993; Korich et al., 2000) was adopted to more precisely quantify oocyst inactivation by various treatments. In this method, the proportion of mice that become infected when orally inoculated with serial doses of oocysts is transformed to a logit, which has a linear relationship to the log[oocyst dose]. Because the precision of the method is critically dependent on the number of mice infected with each oocyst dose, neonatal mice are the natural choice, as one litter of neonatal mice can be used for each dose. To exclude artifacts associated with difference in age between litters, mouse pups are randomly assigned to each treatment.
E.
Animal Models versus Cell Culture
As described earlier, rodent models have a long history of successful application to testing C. parvum oocyst infectivity in the context of the development of therapeutic agents and water disinfection methods. The neonatal mouse model, in particular, has been popular because of its sensitivity (Finch et al., 1993), its relative ease of manipulation, and the feasibility of using large number of animals. However, animal models remain poorly suited for high-throughput applications and for routine monitoring of oocyst infectivity in water-testing facilities. The water industry, in particular, has been interested in simple in vitro alternatives for testing the infectivity of waterborne oocysts. Research was performed by several groups to assess whether in vitro models are valid surrogates for the mouse model. A comprehensive study of the ability of C. parvum to multiply in different transformed cell lines (Upton et al., 1994) indicated that, in spite of its shortcomings, epithelial cell monolayers could be used instead of mice. The sensitivity of neonatal CD-1 mice was compared to that of three cell lines using dose-response experiments (Rochelle et al., 2002). As is typically the case with such experiments, the range of ID50 values obtained in neonatal mice in HCT-8 and CaCo-2 cell monolayers were quite large, but no difference in sensitivity between in vivo and in vitro systems was found. Comparable results were obtained by Joachim et al. (2003), who tested the infectivity of C. parvum oocysts exposed to a disinfectant (25% chlorocresole) in HCT-8 cell monolayers and in neonatal mice (Joachim et al., 2003). Although the experiment was not designed to generate quantitative results, immunofluorescent staining of HCT-8 monolayers and examination of neonatal mice for fecal oocyst shedding detected infectious oocysts following disinfection. Parallel mouse and cell culture assays were also performed to detect infectivity in aged samples of C. parvum oocysts (Jenkins et al., 2003). The neonatal mouse and HCT-8 cell cultures gave similar results in showing diminished infectivity after 6 months of oocyst storage at 15˚C. These results support the view that cell culture can replace the neonatal mouse model for certain studies. In addition to practical consideration favoring in vitro assays, the ability to detect infectious C. hominis and C. parvum oocysts with the same assay also favors the in vitro assay over the mouse. As in some parts of the world C. hominis causes a majority of human infection (Thompson et al., 2005), the ability to detect infectious waterborne oocysts of both human-infectious species is an important consideration.
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Animal Models and Cryptosporidium Host Range
The use of more specialized animal models has led to unexpected discoveries that have challenged established views on host specificity of C. parvum, C. hominis, and C. meleagridis. The fact that C. hominis can infect immunosuppressed gerbils (Baishanbo et al., 2005) and ruminants under laboratory conditions (Akiyoshi et al., 2002; Giles et al., 2001) appears to contradict the widely shared view that C. hominis is strictly anthroponotic. The fact that C. hominis has occasionally been observed in other mammals (Ryan et al., 2005; Smith et al., 2005), or as subpopulations in C. parvum isolates (Tanriverdi et al., 2003), raises the possibility that the host range of this species is not limited to humans. Curiously, C. meleagridis, an avian parasite, has been isolated from humans (Xiao et al., 2001) and experimentally infects a range of mammals including calves and rodents (Huang et al., 2003; Sreter et al., 2000). In light of these findings, the intuitive view that host specificity is determined solely, or primarily, by receptor–ligand interaction may need reassessment. The discovery that some Cryptosporidium species have an extended host range raises interesting questions, and challenges the view that adaptation to different host species drives speciation in this genus (Xiao et al., 2002). This model may only apply to certain species or may be a transient phenomenon in evolution, capable of being supplanted by selective mechanisms favoring a broader host range.
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Tzipori, S., Smith, M., Makin, T., and Halpin, C. 1982a. Enterocolitis in piglets caused by Cryptosporidium sp. purified from calf faeces. Vet. Parasitol. 11, 121–126. Tzipori, S. 1983. Cryptosporidiosis in animals and humans. Microbiol. Rev. 47, 84–96. Tzipori, S. 1988. Cryptosporidiosis in perspective. Adv. Parasitol. 27, 63–129. Tzipori, S., Rand, W., Griffiths, J., Widmer, G., and Crabb, J. 1994. Evaluation of an animal model system for cryptosporidiosis: therapeutic efficacy of paromomycin and hyperimmune bovine colostrumimmunoglobulin. Clin Diagn Lab. Immunol. 1, 450–463. Tzipori, S., Rand, W., and Theodos, C. 1995. Evaluation of a two-phase scid mouse model preconditioned with anti-interferon-gamma monoclonal antibody for drug testing against Cryptosporidium parvum. J. Infect. Dis. 172, 1160–1164. Tzipori, S. 1998. Cryptosporidiosis: laboratory investigations and chemotherapy. Adv. Parasitol. 40, 187–221. Tzipori, S., and Griffiths, J.K. 1998. Natural history and biology of Cryptosporidium parvum. Adv. Parasitol. 40, 5–36. Tzipori, S.R., Campbell, I., and Angus, K.W. 1982b. The therapeutic effect of 16 antimicrobial agents on Cryptosporidium infection in mice. Aust. J. Exp. Biol. Med. Sci. 60, 187–190. Ungar, B.L., Burris, J.A., Quinn, C.A., and Finkelman, F.D. 1990. New mouse models for chronic Cryptosporidium infection in immunodeficient hosts. Infect. Immun. 58, 961–969. Ungar, B.L., Kao, T.C., Burris, J.A., and Finkelman, F.D. 1991. Cryptosporidium infection in an adult mouse model. Independent roles for IFN-gamma and CD4+ T lymphocytes in protective immunity. J. Immunol. 147, 1014–1022. Upton, S.J., Tilley, M., and Brillhart, D.B. 1994. Comparative development of Cryptosporidium parvum (Apicomplexa) in 11 continuous host cell lines. FEMS Microbiol. Lett. 118, 233–236. Vanopdenbosch, E., and Wellemans, G. 1985. Detection of antibodies to Cryptosporidium by indirect immunofluorescence (drop method)—prevalence of antibodies in different animal species. Vlaams Diergeneeskundig Tijdschrift 54, 49–54. Vitovec, J., and Koudela, B. 1992. Pathogenesis of intestinal cryptosporidiosis in conventional and gnotobiotic piglets. Vet. Parasitol. 43, 25–36. Waldman, E., Tzipori, S., and Forsyth, J.R. 1986. Separation of Cryptosporidium species oocysts from feces by using a percoll discontinuous density gradient. J. Clin. Microbiol. 23, 199–200. Widmer, G., Tzipori, S., Fichtenbaum, C.J., and Griffiths, J.K. 1998. Genotypic and phenotypic characterization of Cryptosporidium parvum isolates from people with AIDS. J. Infect. Dis. 178, 834–840. Widmer, G., Akiyoshi, D., Buckholt, M.A., Feng, X., Rich, S.M., Deary, K.M., Bowman, C.A., Xu, P., Wang, Y., Wang, X., Buck, G.A., and Tzipori, S. 2000. Animal propagation and genomic survey of a genotype 1 isolate of Cryptosporidium parvum. Mol. Biochem. Parasitol. 108, 187–197. Xiao, L., Bern, C., Limor, J., Sulaiman, I., Roberts, J., Checkley, W., Cabrera, L., Gilman, R.H., and Lal, A.A. 2001. Identification of 5 types of Cryptosporidium parasites in children in Lima, Peru. J. Infect. Dis. 183, 492–497. Xiao, L., Sulaiman, I.M., Ryan, U.M., Zhou, L., Atwill, E.R., Tischler, M.L., Zhang, X., Fayer, R., and 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. Xu, P., Widmer, G., Wang, Y., Ozaki, L.S., Alves, J.M., Serrano, M.G., Puiu, D., Manque, P., Akiyoshi, D., Mackey, A.J., Pearson, W.R., Dear, P.H., Bankier, A.T., Peterson, D.L., Abrahamsen, M.S., Kapur, V., Tzipori, S., and Buck, G.A. 2004. The genome of Cryptosporidium hominis. Nature 431, 1107–1112. Yang, S., and Healey, M.C. 1994. Development of patent gut infections in immunosuppressed adult C57BL/6N mice following intravenous inoculations of Cryptosporidium parvum oocysts. J. Eukaryot. Microbiol. 41, 67S. Yang, S., Healey, M.C., and Du, C. 1996. Infectivity of preserved Cryptosporidium parvum oocysts for immunosuppressed adult mice. FEMS Immunol. Med. Microbiol. 13, 141–145.
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20 In Vitro Cultivation*
Michael J. Arrowood
CONTENTS I. II. III.
Introduction .................................................................................................................................. 500 Background and Chronology ....................................................................................................... 500 Parasite Production and Purification............................................................................................ 505 A. Oocyst Sources and Storage Conditions......................................................................... 505 1. Oocyst Sources................................................................................................. 505 2. Oocyst Storage Conditions .............................................................................. 506 3. Oocyst Production in Mice.............................................................................. 506 4. Oocyst Production in Neonatal Calves............................................................ 507 B. Stool Collection............................................................................................................... 507 C. Oocyst Purification.......................................................................................................... 507 1. Removal of Fecal Fat and Mucus.................................................................... 507 2. Stool Sieving .................................................................................................... 508 3. Sucrose Gradients ............................................................................................ 509 4. Cesium Chloride Gradients.............................................................................. 509 5. Alternate Methods............................................................................................ 509 D. Excystation of Sporozoites ............................................................................................. 510 E. Sporozoite Purification.................................................................................................... 510 1. Sieving Technique ............................................................................................ 510 2. DE-52 Anion Exchange Chromatography....................................................... 511 3. Percoll® Gradients............................................................................................ 511 4. Sporozoite Viability and Infectivity................................................................. 511 IV. In Vitro Culture............................................................................................................................. 511 A. Oocyst and Sporozoite Preparation ................................................................................ 511 1. Oocyst Age....................................................................................................... 511 2. Preinoculation Labeling of Oocysts and Debris with Fluorescent Dye ......... 512 3. Taurocholate Treatment.................................................................................... 514 4. Inoculum Density............................................................................................. 514 B. Host Cells and Culture Environment.............................................................................. 515 1. Host Cell Selection .......................................................................................... 515 2. Atmosphere Selection ...................................................................................... 515 3. Culture Media .................................................................................................. 515 C. Quantification of Developing Parasites in Culture......................................................... 516 1. Microscopic Enumeration................................................................................ 516 2. Enzyme Immunoassays (EIA) ......................................................................... 518 3. Molecular Methods and Other Alternatives .................................................... 518 D. Culture in Avian Embryos .............................................................................................. 520 V. Concluding Comments................................................................................................................. 520 References........................................................................................................................... 520
*
The findings and conclusions in this report are those of the author and do not necessarily represent the views of the Centers for Disease Control and Prevention.
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I.
Cryptosporidium and Cryptosporidiosis, Second Edition
Introduction
For over 20 years, attempts have been made to cultivate Cryptosporidium species in vitro, and significant progress has been made in establishing routine methods suitable for many laboratory studies. Building on the groundwork of standard cell culture of mammalian host cells and successful cultivation of various coccidian parasites (Doran, 1973, 1982; Speer, 1983), in vitro culture of Cryptosporidium parvum through asexual and sexual stages has been accomplished. Although yet controversial, in vitro cultivation in the absence of host cells has been reported (Hijjawi et al., 2004). Many studies regarding in vitro cultivation have been descriptive and have introduced cell lines with various capacities to support cryptosporidial development. Methods for parasite preparation, inoculation, direct enumeration or indirect quantification, and more specific studies evaluating culture media composition and atmospheric conditions conducive to support Cryptosporidium growth have been especially valuable.
II.
Background and Chronology
Key publications that introduced particular cell lines or models and modifications that encouraged the adoption of useful Cryptosporidium culture systems are summarized in the following text (and in Table 20.1). The first successful in vitro culture of Cryptosporidium asexual life-cycle stages was described by Woodmansee and Pohlenz (1983). A bovine oocyst isolate stored in 2% aqueous potassium dichromate (K2Cr2O7), then purified by repeated sucrose flotation (1.18 g/cm3, 1200 × g, 5 min), and preincubated in a reducing environment (0.02-M cysteine hydrochloride in 0.85% sodium chloride and 50% CO2 in air) was suspended in an excystation fluid (0.75% sodium taurocholate and 0.25% trypsin in phosphatebuffered saline [PBS]) for 30 min before inoculation onto host cells. Human rectal tumor (HRT) cells maintained in RPMI 1640 supplemented with 10% heat-inactivated fetal bovine serum (FBS) at 37°C and 5% CO2 were incubated for 48 h in Leighton tubes before oocyst/sporozoite inoculation. Brightfield and electron microscopy revealed asexual stages, but no sexual stages through 3 days post inoculation. Indirect immunofluorescence using immune human sera revealed distinctly reactive life-cycle stages. Endoderm cells of the chorioallantoic membrane (CAM) of chicken embryos supported development of Cryptosporidium from humans and calves (Current and Long, 1983). There were no morphologic differences in the developmental stages of the human and calf isolates grown on the CAM, and oocysts collected from the CAM were infectious for suckling mice. Despite this initial report of success in cultivating human and calf Cryptosporidium isolates, no reports completely replicate the results. In 1984, human fetal lung cells (HFL), primary chicken kidney (PCK), and porcine kidney cells (PK10) supported development of a human Cryptosporidium isolate (Current and Haynes, 1984). Although all three cell lines supported development of the entire life cycle, HFL cells produced the greatest yield. Because oocysts were derived from an AIDS patient or from goats inoculated with the same isolate, it was probably C. parvum (zoonotic), as opposed to C. hominis, which is essentially restricted to humans (anthroponotic). Oocysts were stored in 2.5% K2Cr2O7 at 4°C, purified by Sheather’s flotation, washed with PBS, incubated with antibiotics at 37°C (penicillin, streptomycin, and amphotericin B), and excysted for 30 to 60 min at 37°C in PBS containing 0.75% sodium taurocholate and 0.25% trypsin. The excysted mixture of oocysts, oocyst walls, and free sporozoites was washed with PBS, suspended in growth medium, and inoculated onto cells in Leighton tubes at approximately 1.5 × 104 sporozoites/tube (5 cm2). Culture medium consisted of minimal essential medium (MEM) with Earle’s salts, 10% FBS, L-glutamine, penicillin (100 IU/mL), streptomycin (100 μg/mL), and amphotericin B (0.25 μg/mL). Preparations were maintained at 37°C in 5% CO2. Type I meronts observed at 96 h, were interpreted as evidence for asexual stage recycling in culture. Other laboratories were inspired to pursue HFL cells for Cryptosporidium cultivation, but anecdotal reports suggest that attempts to replicate the success of the original study were disappointing.
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TABLE 20.1 Cryptosporidium: Selected Chronology of In Vitro Cultivation Efforts
Cell Linea
Oocyst Source
Cryptosporidium Life-Cycle Stage Development Asexual Sexual Oocysts
Reference
HRT
Bovine
Yes
ND
ND
Chicken CAM
HFL
Human, bovine Human; caprine-passaged Chicken (C. baileyi) Human; caprine-passaged
Yes Yes Yes Yes
Yes NR Yes Yes
Yes NR Yes Yes
PCK, PK-10
Human; caprine-passaged
Yes
Yes
Yes
BHK (CRL-1632) MDCK (CCL-34)
Human; caprine-passaged
Yes
NR
NR
Woodmansee and Pohlenz, 1983 Current and Long, 1983 Naciri et al., 1986 Lindsay et al., 1988 Current and Haynes, 1984 Current and Haynes, 1984 Naciri et al., 1986
Caprine Bovine Bovine Bovine Human Human, bovine Human Human, bovine
Yes Yes Yes Yes ? Yes Yes Yes
NR Yes Yes Yes ? NR Yes Yes
NR Yesb Yes Yesb Yes NR Yes NR
Lumb et al., 1988 Gut et al., 1991 Rosales et al., 1993 Arrowood et al., 1994 Datry et al., 1989 Griffiths et al., 1994 Datry et al., 1989 McDonald et al., 1990
Human, bovine Human
Yes Yes
NR ?
NR NR
Bonnin et al., 1990 Flanigan et al., 1991
Bovine
Yes
NR
NR
Kuhls et al., 1991
Bovine
Yes
Yes
?
Rasmussen et al., 1993
Bovine
Yes
Yes
NR
Adams et al., 1994
Bovine Bovine Bovine Bovine Bovine
Yes Yes Yes Yes Yes
Yes Yes Yes Yes Yes
Yes Yes Yes Yes Yes
Human, bovine
Yes
Yes
Yes
Human; bovine- and caprine-passaged Bovine
Yes
Yes
Yes
Yang et al., 1996 Gut et al., 1991 Upton et al., 1994c Villacorta et al., 1996 Upton et al., 1994b Upton et al., 1994a Upton et al., 1995a Upton, 1997 Nesterenko, 1997 Hijjawi, 2001 Lawton et al., 1997
Yes
Yes
Yes
Deng and Cliver, 1998
NR
NR
NR
Dawson et al., 2004
Yes
Yes
Yes
Hijjawi et al., 2004
Caco-2 (HTB-37) Primary rat hepatocyte L929 (CCL-1) LGA HT29.74 (HTB-38) Intestine 407 (CCL-6) RL95-2 (CRL-1671) T84 (CCL-248) BFTE MDBK (CCL-22) HCT-8 (CCL-244)
THP-1 (ECACC 88081201) BS-C-1 (ATCC CCL26) MRC-5 (ECACC 97112601) Axenic (no host cells)
Human (C. hominis); Cervine; caprinepassaged Bovine; murine-passaged
Note: NR = not reported; ND = not detected; ? = not clear from manuscript description. a
b
Cell line name or abbreviation (and where available the ATCC reference number in parentheses). (See Table 20.3 for descriptions. See text for culture details.) = Immature and possible mature forms.
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Baby hamster kidney (BHK) cells and chicken embryos supported Cryptosporidium growth (Naciri et al., 1986). The isolate, obtained from an immunocompromised patient and propagated in goats, was identified as C. muris but was probably a zoonotic strain of C. parvum. After 45 passages at weekly intervals, there were mostly free sporozoites and merozoites in the chorioallantoic fluid. Chicken embryos supported growth of C. baileyi (AU-B1 isolate) (Lindsay et al., 1988). Oocyst-laden chicken feces were sieved, stored in 2.5% K2Cr2O7 at 4°C, and oocysts were purified by Sheather’s flotation, washed with distilled water, and stored in Hanks’ balanced salts solution (HBSS) at 4°C until “sanitized” in 0.26% sodium hypochlorite. Residual hypochlorite was removed by washing with HBSS before oocyst inoculation into chicken embryos (4 × 105 oocysts/embryo). Embryos were incubated at 40°C up to 6 days. Embryo-generated oocysts were treated with sodium hypochlorite (0.5%, 4°C, 5 min), stored in HBSS (supplemented with 100 IU/mL penicillin and 100 μg/mL streptomycin) at 4°C, and used to serially propagate the isolate through 20 embryo passages. Efforts to cultivate the C. baileyi isolate in vitro failed in multiple avian and mammalian primary and continuously growing cell lines (16 total). Attachment and invasion of Madin–Darby canine kidney (MDCK) cells by sporozoites from Cryptosporidium isolated from a naturally infected goat kid was reported by Lumb et al. (1988). Diarrheic feces were stored in 1% K2Cr2O7 at 4°C, treated with Sputolysin® (Boehringer Ingelheim, Ridgefield, CT), subjected to ether sedimentation, and purified using discontinuous Ficoll® (Pharmacia, Piscataway, NJ)/sodium diatrizoate gradients. MDCK cells were cultured in DMEM medium supplemented with 10% FBS and HEPES buffer. Electron microscopy revealed numerous asexual stages (meronts) at 48 h post infection. Human colonic adenocarcinoma (Caco-2) cells and primary rat hepatocyte cells supported complete life-cycle development of Cryptosporidium from an AIDS patient (Datry et al., 1989). Stools were collected and stored in 2.5% K2Cr2O7. Oocysts were purified via Sheather’s flotation, washed with distilled water, “sanitized” with 1.75% sodium hypochlorite for 1 h, excysted in trypsin/EDTA/taurocholate at 37°C for 1 h, washed, suspended in DMEM or RPMI, and 1 × 105 sporozoites/mL were inoculated onto Caco-2 cells (DMEM, 20% FBS) and primary rat hepatocytes (MEM, 10% FBS). Asexual, sexual, and oocyst stages were identified in hepatocyte cultures. Unidentified stages were observed in Caco-2 cultures; only oocysts were clearly identified. Mouse L929 fibroblasts were used to test the efficacy of antimicrobial agents against development of bovine and human isolates of Cryptosporidium (McDonald et al., 1990). Oocyst-laden stools were diluted with water, sieved, and defatted when necessary by ether sedimentation. Sediment was removed by flotation on saturated sodium chloride or Sheather’s sugar. Recovered oocysts were washed with water and stored in 2.5% K2Cr2O7 at 4°C. Purified oocysts were treated with 0.5% trypsin in HBSS at 37°C for 30 min, excysted in 0.4% sodium taurocholate at 37°C for 45 min, washed with RPMI 1640 culture medium, and inoculated onto cells at 1 × 105 oocysts/culture chamber (24-well plate well or well of 8chambered slide). Cells were incubated for 24 h at 37°C in candle jars, washed with HBSS, fixed with methanol, and Giemsa-stained. Asexual stages predominated, with less than 10% of sexual stages apparent at 72 h. Parasite numbers peaked between 48 and 72 h, and no evidence of stage recycling was observed. LGA cells (small intestine carcinoma from Lewis rats) were used to obtain development of asexual stages of Cryptosporidium of bovine and human origin (Bonnin et al., 1990). Feces were stored in 2.5% K2Cr2O7, at 4°C. Oocysts were purified via flotation on saturated sodium chloride, “sanitized” with sodium hypochlorite, excysted in 0.04% sodium taurocholate in Ham’s F10 medium, washed, and inoculated onto cells at 2 × 104 sporozoites/0.5 cm2. Developing stages were visualized after methanol fixation and Giemsa staining or by transmission electron microscopy. Differentiated and undifferentiated HT29.74 human colon adenocarcinoma cells were compared for their support of C. parvum growth (Flanigan et al., 1991). Oocysts from humans with AIDS were stored in 2.5% K2Cr2O7 at 4°C, sieved (40-gauge stainless steel), purified over saturated sodium chloride, sieved through nylon mesh (30 µm), “sanitized” with 0.05% sodium hypochlorite (10 min), suspended in antibiotic mixture (100 IU/mL penicillin, 0.1 mg/mL streptomycin, 0.2 mg/mL neomycin) and stored at 4°C before inoculation onto cells. Undifferentiated cells were maintained in RPMI 1640, 10% dialyzed FBS, 24-mM sodium bicarbonate, 1-mM sodium pyruvate, 20-mM HEPES, 2-mM L-glutamine,
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penicillin, and streptomycin. Differentiated cells were maintained in Leibovitz L-15 medium, 10% FBS, 5-mM galactose, 6-mM pyruvate, 1-mM L-glutamine, 20-mM HEPES, penicillin, and streptomycin. Oocysts were inoculated at 105 to 106/cm2 onto cells. Cultures grown in slide chambers were fixed and stained with hematoxylin and eosin and examined by bright-field microscopy. Differentiated cells supported fivefold higher levels of development than undifferentiated cells. The data suggest the infection efficiency was low and development was primarily restricted to asexual stages through 72 h post infection. This model showed the importance of microtubules in host cell invasion and parasite development, because treatment with either colchicine or vinblastine significantly reduced parasite numbers in a dosedependent manner (Wiest et al., 1993). Of 15 cell lines compared for potential support of C. parvum growth in vitro, 3 cell lines supported no growth, 11 cell lines supported poor development (less than 0.1% infection), but MDCK and MDBK cells became heavily infected (up to 90%) (Gut et al., 1991). Oocysts of C. parvum from a bovine source (AUCP-1) were sieved, purified by sucrose density gradient centrifugation, “sanitized” with undiluted commercial bleach for 5 min at 4°C, washed with RPMI 1640 medium, and stored at 4°C until used. Sporozoites, excysted by suspending oocysts in 0.75% sodium taurocholate under reduced oxygen, were passed through 3-µm polycarbonate filters, suspended in culture medium, and inoculated onto host cells. The optimal culture medium for MDCK cells was RPMI 1640, 10% FBS, 2.5 g/L sodium bicarbonate, 1-mM sodium pyruvate, 2-mM L-glutamine, 100 IU/mL penicillin, 100 μg/mL streptomycin, 25-mM HEPES, nonessential amino acids, 0.24 U/mL insulin, 13.6 mg/L hypoxanthine, 2.9 mg/L thymidine, 50-µM 2-mercaptoethanol, 0.1% yeast extract, 1% Ficoll 400, 4.5 g/L glucose, FBS (reduced to 1% during parasite development), and 0.1% rabbit bile. Development through sexual stages, including immature oocysts (partial oocyst wall formation) was observed. Although earlier publications were encouraging, the success reported in this study provided the impetus for several other laboratories to pursue in vitro culture of Cryptosporidium more aggressively. The utility of this model for evaluating antibody-mediated life-cycle modulation was reported by Doyle et al. (1993). Additional studies demonstrated the development of C. parvum in MDBK cells (Upton et al., 1994c; Villacorta et al., 1996). Human embryonic intestine 407 cells (ATCC-CCL-6) supported development of asexual stages of a C. parvum bovine isolate and treatment with carbohydrates and/or lectins influenced the outcome of infection (Kuhls et al., 1991). Human endometrial carcinoma cell line RL95-2 supported optimal development of a C. parvum bovine isolate when cells were cultured for 7 days before parasite inoculation onto coverglass disks (Rasmussen et al., 1993). Oocysts were treated with 2% sodium hypochlorite, washed five times with PBS, excysted at 37°C in 0.75% sodium taurocholate and 0.25% trypsin (in PBS), washed with PBS, suspended in culture medium, filtered through 3-µm polycarbonate filters, and inoculated onto host cells in a 1:1 mixture of Dulbecco’s modified Eagle’s medium and Ham’s F12 medium, supplemented with 10% FBS, 2 g/L sodium bicarbonate, 10-mM HEPES, 5-μg bovine insulin, 100 IU/mL penicillin G, 100 μg/mL streptomycin, and 0.25 μg/mL amphotericin B. After variable culture periods, cells on coverglass disks were washed in phosphate buffer, fixed with Bouin’s fixative for 2 h, washed with ethanol, and stained with Giemsa stain. Apparent asexual and sexual stages were observed by bright-field microscopy, and possible oocysts via scanning electron microscopy. MDCK cells supported development of a bovine isolate of C. parvum better than Vero, HeLa, and McCoy cells (comparative data were not provided) (Rosales et al., 1993). MDCK cells were cultured under 5% CO2 at 37°C in MEM supplemented with 10% FBS. Excysted sporozoites in MEM without FBS were inoculated onto 12 to 24 h nonconfluent MDCK cells on glass coverslips. Inoculum was replaced with MEM plus FBS for periods up to 120 h. MDCK cells grew naturally polarized directly on coverslips in culture plates or on permeable membrane supports. Cultures on glass coverslips were removed, fixed with acetone (20°C), and stained with Alcian blue and Giemsa stains. All life-cycle stages were seen by bright-field microscopy, including asexual stages persisting through 120 h post inoculation, suggesting asexual stage recycling. Researchers from this laboratory described development of a C. parvum bovine isolate in unsensitized mouse peritoneal macrophages, emphasizing the ability of this parasite to infect nonepithelial cells (Martinez et al., 1992). Its persistence in macrophages was interpreted as evidence of its escape from lysosomal degradation.
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A modified MDCK culture system was developed for use in evaluating potential anticryptosporidial agents and to study life-cycle stage development of a C. parvum bovine-derived isolate (IOWA) (Arrowood et al., 1994). MDCK and Caco-2 cells were grown under 5% CO2 at 37°C in conventional FBSsupplemented high-glucose DMEM culture medium and compared with MDCK cells grown in a synthetic, serum-free culture medium (Ultraculture®, Cambrex Bio Science, Walkersville, MD). Cryptosporidial development in the MDCK/Ultraculture®-based model outperformed MDCK (2 to 5 fold) and Caco-2 (2 to 20 fold) models using serum-supplemented DMEM media. Detailed culture conditions are presented in the following text. The model has been used to evaluate potential anticryptosporidial agents and for disinfection studies. (Arrowood et al., 1994, 1996; Shin, 2001; You et al., 1996a,b, 1998; Venczel et al., 1997). Colonic epithelial cells (T84) were used to examine the role of bovine C. parvum infection on host cell membrane integrity and the modulation of macromolecular flow (Adams et al., 1994). T84 cells were grown under 5% CO2 at 37°C (preincubated for 7 days to facilitate polarization and differentiation) in DMEM and Ham’s F12 media (1:1) supplemented with 5% fetal calf serum (FCS), 14-mM sodium bicarbonate, 15-mM HEPES, penicillin, and streptomycin. Infection was achieved with 105 oocysts and alteration of epithelial barrier function was correlated with oocyst dose. The parasite was found responsible for disrupting the epithelial cell barrier, not simply opening transcellular channels for ion flow. Asexual and sexual stages were identified in cultures by light and electron microscopy. In a related study, Caco-2 cells were grown on permeable membrane supports (Griffiths et al., 1994). Isolates of human- and bovine-derived C. parvum were inoculated onto Caco-2 and Caco-2a cells grown under 5% CO2 at 37°C in DMEM supplemented with 20% FBS, and 3.5g/L glucose. Transmonolayer resistance (as a measure of membrane integrity and permeability) declined in a dose- and time-dependent manner following parasite inoculation. Molecules less than 1 kDa leaked through the monolayers, but proteins 1.8 kDa or larger were excluded. Cellular lactate dehydrogenase was released into culture medium on the apical side of the monolayer where parasites were located, but not on the basal side. Propidium iodide uptake by infected cells was interpreted as evidence that cryptosporidial infection killed host cells. Results did not achieve significance until data from both human and bovine isolates were pooled (either data set alone was not significant). Primary bovine fallopian tube epithelial (BFTE) cells were efficient hosts for a C. parvum bovine isolate (Yang et al., 1996). Giemsa-stained cells grown on coverslips were examined by bright-field microscopy or scanning and transmission electron microscopy. The authors concluded that monolayers contained de novo mature, infectious oocysts because ethanol treatment of the monolayer did not prevent infection of mice inoculated with infected monolayers, and stages other than oocysts were presumably killed by the ethanol. For the lengthy description of host cell isolation, establishment in culture, and electron microscopy methods, the reader is directed to the original publication. Several studies examined the role of atmospheric conditions and media composition on the performance of various cell lines supporting C. parvum in vitro development (Upton et al., 1991, 1994a–c, 1995; Eggleston et al., 1994; Woods et al., 1995; Nesterenko et al., 1997; Upton, 1997). Ultimately, human colonic tumor (HCT-8) cells performed best for these investigators when cultivated under 5% CO2 at 37°C, in RPMI 1640 basal medium supplemented with 2-mM L-glutamine, 15-mM HEPES buffer, 50-mM glucose, 35 μg/mL ascorbic acid, 4.0 μg/mL para-aminobenzoic acid, 2.0 μg/mL calcium pantothenate, and 1.0 μg/mL folic acid. The medium was filter-sterilized after adjusting the pH to 7.4 and 10% FBS added. When needed, recommended antibiotic supplements were 100 U/mL penicillin, 100 μg/mL streptomycin, and 0.25 μg/mL amphotericin B. The nonadherent human monocytic cell line THP-1 (ECACC 88081201) was found permissive for a human isolate of C. parvum passaged through calves or goat kids (Lawton et al., 1997). THP-1 cells were grown at 37°C in a CO2 incubator using RPMI-1640 basal medium supplemented with 25-mM HEPES, 10% fetal calf serum, penicillin (200 IU/mL), and streptomycin (200 μg/mL). Sucrose gradient purified oocysts were “sanitized” using 1.25% sodium hypochlorite (5 min, 0°C), inoculated into THP1 cell suspension cultures (1:1 to 1:2 ratio), incubated for 2 to 6 h in a candle jar (to promote excystation), and removed from cultures by low speed centrifugation (160 × g, 4 min). Microscopic examination of Giemsa-stained and antibody-labeled (immunofluorescence) cultures showed development of parasites,
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including asexual and sexual stages. Despite the apparent identification of the entire life cycle, including oocysts, no successful subculture was accomplished. The African green monkey kidney cell line BS-C-1 supported development of the complete life cycle of a C. parvum bovine isolate (Deng and Cliver, 1998). Cells were grown in Eagle’s MEM supplemented with 2.0-mM L-glutamine and 5% FBS. Oocysts were purified using sucrose and cesium chloride gradients, treated with a 1/10 dilution of commercial bleach for 10 min at 0°C, inoculated onto cultures, and incubated for 3 h in a candle jar to promote excystation and invasion of sporozoites. Cultures were washed with PBS, culture medium replaced, and incubated at 37°C in a 5% CO2 incubator. Direct oocyst inoculation performed better than inoculation with purified sporozoites. Secondary human lung fibroblast cells (MRC-5 cells, ECACC, Salisbury) supported development of C. parvum and C. hominis and were used to evaluate oocyst survival under conditions mimicking various food environments (Dawson et al., 2004). Oocysts from human stools (C. hominis) were purified over saturated NaCl flotation. Oocysts of C. parvum originally isolated from deer and passaged in lambs were obtained from a commercial source. MRC-5 cells were grown in Medium 199 containing Earle’s salts, essential amino acids, and L-glutamine and was supplemented with 0.075% sodium bicarbonate, 10% FCS, 3 mg/mL sodium benzylpenicillin, and 160 μg/mL gentamycin. When oocysts were inoculated, the medium also contained 20 μg/mL amphotericin B. Oocysts were not treated with bleach or excystation solutions before inoculation into cell cultures. After incubation for 5 days at 37°C in a 5% CO2 incubator, cultures were fixed with acetone, stained with auramine-phenol, and examined with the aid of a fluorescence microscope. Fluorescing structures in 5 or 10 randomly selected microscopic fields (200 or 400× total magnification) were counted and infectivity estimated. No microscopic images were provided in the publication, and no specific life-cycle stages were identified. A novel extracellular life-cycle stage was described for C. andersoni grown in HCT-8 cells (Hijjawi et al., 2002). A follow-up report described development of C. parvum in vitro in the absence of host cells (Hijjawi et al., 2004). The C. parvum bovine isolate was passaged through mice, and then purified from gut homogenates using ether extraction and Ficoll® gradients. It was excysted in acidic trypsin solution, and inoculated into culture flasks containing a semisolid base of coagulated newborn calf sera and an RPMI 1640-based maintenance medium adjusted to pH 7.4 containing 0.03-g L-glutamine, 0.3g sodium bicarbonate, 0.02-g bovine bile, 0.1-g glucose, 25-mg folic acid, 100-mg 4-aminobenzoic acid, 50-mg calcium pantothenate, 875-mg ascorbic acid, 1% FCS, 0.36-g HEPES buffer, 10,000-IU penicillin G, and 0.01-g streptomycin (per 100 mL medium). Structures, including pigmented forms, found in the culture flasks at various times post inoculation were described as extracellular stages of Cryptosporidium, and an “extracellular life cycle” was proposed. The results are conflicting; some studies provide support and others fail to reproduce the results. Gregarine-like extracellular structures consistent with those described by Hijjawi and colleagues were observed by Rosales et al. (2005), but efforts to reproduce host-cell-free development of C. parvum failed in a study by Girouard et al. (2006). Other attempts to reproduce the results have also failed (Arrowood, unpublished). It is not clear if the inconsistent results reflect isolate-specific effects. Direct comparisons of the isolate used by Hijjawi and colleagues and the other C. parvum isolates might yield important clues about the potential “axenic” development of Cryptosporidium.
III. Parasite Production and Purification A. 1.
Oocyst Sources and Storage Conditions Oocyst Sources
Cryptosporidium oocysts for in vitro cultivation studies have primarily come from clinical human or veterinary stool samples or from oocysts generated in experimentally infected laboratory animals or livestock (see Chapter 19). Zoonotic C. parvum oocysts can be generated in low-to-moderate numbers in rodent animal models, primarily neonatal, immunocompromised, or immunosuppressed mice or rats. Large-scale production is generally accomplished in neonatal livestock, especially bovine calves, goat
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kids, or lambs (Arrowood and Sterling, 1987; Current, 1990; Arrowood and Donaldson, 1996; Upton, 1997). Although no cloned stocks of C. parvum isolates are available, oocysts can be obtained from several research or commercial sources. Clinical isolates are sometimes useful for research studies, but are often more difficult to purify and typically have shorter shelf lives than routinely passaged isolates. A review of in vitro cultivation publications indicates that a majority of studies employed C. parvum or C. parvumlike oocysts (Table 20.1). The use of other strains or species has been limited, but in general such studies employed oocysts derived from clinical isolates. Inoculating MDCK, Caco-2, and HCT-8 cells with oocysts of C. parvum (zoonotic) and C. hominis (anthroponotic) resulted in successful development through sexual life-cycle stages (Arrowood et al., 1994). Although partial-to-complete oocyst wall formation was evident in these cultures, significant development of mature oocysts was not observed. Short- and long-term growth of zoonotic and anthroponotic isolates was reported in HCT-8 cells (Hijjawi et al., 2001). Oocyst yields from in vitro culture are insufficient for most laboratory studies; consequently, experimentally infected animals continue to be the primary source for research purposes. Cryptosporidium hominis has been propagated in gnotobiotic piglets (Widmer et al., 2000; Akiyoshi et al., 2002), but this model is not routinely available to most researchers. It has recently been propagated in an immunosuppressed hamster model that might be accessible to more investigators (Chapter 19).
2.
Oocyst Storage Conditions
Clinically or experimentally derived oocysts are preferentially stored at 4°C in 2.5% aqueous potassium dichromate (K2Cr2O7) as this oxidizer inactivates other microbial contaminants and allows oocysts to survive for extended periods ( 6 to 12 months). K2Cr2O7 also “neutralizes” most of the offensive odor of feces, making processing much less demanding. Storage in crude stool suspensions versus purified oocyst suspensions is largely a matter of laboratory preference. Because many purification methods preferentially recover intact, viable oocysts, it is recommended to batch-purify oocysts soon after collection and store them in small volumes of storage solutions. Because K2Cr2O7 must be disposed of as a hazardous waste, it is less costly to use the smallest quantity necessary for processing and storage. If oocysts are preferred without exposure to dichromate, it is recommended that the stool suspension be processed quickly because microbial overgrowth leading to anaerobic conditions will shorten the useful “shelf life” of the oocysts. It is impractical to use antibiotics in minimally processed stool suspensions; antibiotics are typically added after oocyst purification. A typical antibiotic mixture useful in cell culture can be used with cryptosporidial oocysts suspended in saline (0.85%) or PBS (0.01-M, pH 7.2), but it is recommended to double (2×) the normal working concentrations: 100× stock contains 10,000-IU penicillin, 10,000-μg streptomycin, and 25-μg amphotericin B per mL. Alternatively, oocysts may be suspended in PBS supplemented with 1.5× concentration of the following antibiotic mixture: 100× stock contains 5.0-mg gentamycin, 2.0-mg lincomycin, and 10-mg ampicillin per mL (Meloni and Thompson, 1996). Oocyst suspensions should be routinely centrifuged (1500 × g, 10 min), and spent antibiotic suspension aspirated and replaced with fresh antibiotic solutions in saline or PBS. The frequency of replacement is dependent on the purity of the oocyst suspension, but weekly or biweekly replacement is suggested until the oocysts are depleted in laboratory studies. Despite promising studies, no validated methods have been adopted for cryopreserving C. parvum oocysts or sporozoites (Rossi et al., 1990; Fayer et al., 1991). The failure of standard cryopreservation techniques, including the use of dimethyl sulfoxide (DMSO), glycerol, and extracellular matrices (e.g., hydroxyethyl starch) previously reported useful for cryopreserving Plasmodium (Hollingdale et al., 1985), continues to impede the establishment of standardized cryptosporidial isolates or cloned stocks.
3.
Oocyst Production in Mice
A wide range of mice can be infected as neonates with C. parvum (Sherwood et al., 1982; Ernest et al., 1986; Novak and Sterling 1991). Although oocyst production in adult immunocompromised or immunosuppressed rodents has been advocated as a means of generating moderate numbers of oocysts (Rehg et al., 1987; Leitch and He, 1994; Yang and Healey, 1994), neonatal mice are an attractive alternative in terms of cost, husbandry, and simplicity of technique (Meloni and Thompson 1996; Upton, 1997).
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Briefly, neonatal mice (4 to 8 days old) are inoculated with 104 to 105 oocysts by gavage (orogastric intubation using a 24-gauge inoculating needle or equivalent) or simply delivering the inoculum (5 to 10 µL in water, saline, PBS, or other appropriate buffer or culture media) to the back of the mouth using a micropipette and disposable tip. At 7 to 8 days post inoculation, remove the intestine and place in 1to 2-mL PBS or 2.5% K2Cr2O7 and homogenize with a Teflon®-coated or glass tissue grinder. Dilute the homogenate with PBS or K2Cr2O7 and, if necessary, process over sucrose gradients and/or cesium chloride gradients (described in the following text). Oocysts should be stored at 4°C in 2.5% K2Cr2O7. Approximately 500-fold amplification is achieved, typically 2 to 8 × 106 oocysts per mouse (Meloni and Thompson, 1996; Upton, 1997).
4.
Oocyst Production in Neonatal Calves
Large oocyst yields may be obtained from neonatal calves. Because C. parvum is extremely common among dairy herds (the usual source of calves), it is recommended that calves be obtained as soon after birth as possible and that they receive colostrum before oocyst inoculation. Ideally, calves should be approximately 2 days old when inoculated, and the inoculum should be relatively large, between 1 × 107 and 1 × 108 oocysts. The large inoculum and early infection should dominate any “local” isolate picked up by the calf before acquisition for experimental infection. Despite these precautions, isolate competition, and even replacement, have been reported. Valuable isolates may be maintained in specific pathogen-free mice as “seed” stocks if vendors routinely deliver calves already infected before experimental inoculation occurs. Calves should be fed commercial milk replacer (with or without antibiotics), supplemented as necessary with nonfat dry milk and electrolyte solution until day 4 post inoculation, whereupon calves will receive the electrolyte mixture alone. This feeding strategy provides appropriate nutrition, electrolytes, and fluids for scouring calves while reducing the lipid load in the stool (facilitating oocyst purification). Stool collection begins 5 days after inoculation and continues through day 12 for a total of 8 days. Variation in oocyst shedding often leads to discarding stool collected on day 5, and sometimes days 11 and 12 postinoculation. Given the labor-intensive nature of oocyst purification, oocyst levels in the raw or sieved stool collected each day should be estimated before forwarding for purification.
B.
Stool Collection
A number of improvements in the production and purification process for stool-derived oocysts have been introduced to minimize labor, consumption of materials, and generation of hazardous waste (Arrowood and Donaldson, 1996). Stool specimens may be collected unpreserved or diluted with tap water, saline, or suspended in a storage medium composed of aqueous K2Cr2O7 (2.5% final concentration, w/v), often accomplished by mixing equal volumes of liquid stool with 5% w/v aqueous K2Cr2O7. Alternatively, liquid stool samples can be centrifuged (1500 × g, 10 min), supernatant aspirated, and the pellet resuspended in 2.5% K2Cr2O7. Because oocysts are resistant to routine disinfection and remain infectious for extended periods, laboratory manipulation of these agents should be conducted under level 2 biological hazard conditions (Angus et al., 1982; Campbell et al., 1982; Pavlasek, 1984; Blewett, 1989).
C.
Oocyst Purification
Figure 20.1 illustrates a simple stool purification scheme that yields oocysts and sporozoites suitable for in vitro culture studies. Details of the process are described in the following text.
1.
Removal of Fecal Fat and Mucus
Stool specimens with excessive fat content may be defatted with organic solvents before further processing to enhance oocyst recovery and purity (Current, 1990). Replace K2Cr2O7 (if necessary) in stool suspension with saline or tap water. Vigorously mix the stool suspension for ~30 s with diethyl ether or ethyl acetate at a ratio of 2:1 stool to solvent, centrifuge at 1000 × g for 10 min, discard the supernatant
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Fresh stool Mix diarrheic stool with equal volume 5% K2Cr2O7 or formed stools with equal volume 2.5% K 2Cr2O7 at 4°C.
Stool can be stored at 4°C for ≥ . 6 months.
Alternatively, dilute in water or saline and process as follows at 4 C.
Homogenize stool and process sequentially through #20 and #60 mesh sieves (or cheese cloth) to remove large particles.
Adjust supernatant:settled solids ratio to approximately 3:1 to 4:1 (after overnight settling).
Apply stool suspension to discontinuous sucrose gradients, centrifuge. Repeat once Recover oocyst-enriched layer between sucrose layers.
Apply sucrose gradient-purified oocysts to cesium chloride step gradient, centrifuge.
Recover oocyst-enriched layer at sample:gradient interface.
Store at 4°C in 2.5% K2Cr2O7 or in PBS or HBSS supplemented with cell culture antibiotic
FIGURE 20.1 Flowchart summarizing the purification and preparation of Cryptosporidium oocysts for use in cell culture studies. (See Figure 20.3 for example images of gradient techniques.)
(oocysts are in the pellet), and wash the pellet with saline or PBS at least twice to remove residual solvent. Resuspend oocysts in saline or 2.5% K2Cr2O7. Stools with high levels of mucus can benefit from treatment with reducing agents (e.g., Sputolysin®) or potassium hydroxide (0.1%) at room temperature for 10 min, followed by centrifugation (1500 × g, 10 min). Without such treatment, oocysts may not adequately sediment from the stool suspension. Fat removal and mucus dissolution can shorten long-term viability (shelf life) of oocysts, but shortterm viability and infectivity are largely unaffected. Exposure to organic solvents or mucus dissolution techniques may alter the oocyst surface and change oocyst antigens; avoiding these steps may be warranted if antigenic or structural analyses are being performed. For disinfection studies that target the oocyst wall, purification techniques should be selected that minimize changes to oocyst wall integrity.
2.
Stool Sieving
Sieving is applied to stools to remove large debris that interferes with subsequent processing techniques. Stools (“raw” or suspended in tap water, saline, PBS, or K2Cr2O7) are applied sequentially to 20- and 60-mesh stainless steel sieves (850 and 250 µm average porosity, respectively). Use of sieves with higher mesh sizes (lower “pore” diameters) can lead to loss of oocysts. Use saline or tap water and a spatula to assist stool passage through the sieves. Centrifuge at 1500 × g, 10 min, aspirate supernatant, and resuspend pellet in saline or 2.5% K2Cr2O7. A rule of thumb is to maintain a ratio of 3:1 to 4:1 for supernatant to settled solids (after an overnight settling) in preparation for sucrose gradient processing. Overloading gradients or other flotation methods with fecal solids results in lower or “dirtier” yields.
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Sucrose Gradients
Although optimized for neonatal calf feces, the following gradient technique (Figure 20.3) works well with most stool samples and can be modified in a variety of ways for particular laboratory purposes (see the following text). Sieved stool is applied to discontinuous sucrose gradients (scaled according to the available volume of stool) essentially as previously described (Arrowood and Sterling, 1987; Arrowood and Donaldson, 1996). Large-scale gradients are prepared in 200-mL polycarbonate bottles by layering 80 mL of a 1.103-sp. gr. sucrose solution beneath 80 mL of a 1.064-sp. gr. sucrose solution. This “underlaying” technique produces a sharper, less diffuse interface between the sucrose layers. The respective solutions (STP 1:2 and STP 1:4) are prepared by mixing Sheather’s solution (500-g sucrose, 320-mL H2O, 9-mL aqueous phenol (85% w/v)), Tween 80, and PBS as follows: STP 1:2 (sp. gr. 1.103 g/L) = 300-mL Sheather’s solution, 9-mL Tween 80, and 600-mL PBS; STP 1:4 (sp. gr. 1.064 g/L) = 200-mL Sheather’s solution, 9-mL Tween 80, and 800-mL PBS. Forty milliliters of the sieved stool are overlaid onto each sucrose gradient, and the bottle is centrifuged at 1000 × g for 25 min. The top layer and half of the STP 1:4 layer are discarded (approximately 60 mL), and the oocyst-enriched layer occupying the interface between the sucrose layers (approximately 90 to 100 mL) is transferred to a conical 175-mL polycarbonate or polypropylene centrifuge bottle (Nalgene). This fraction contains oocysts. It is diluted to 175 mL with saline, mixed thoroughly, and the bottles centrifuged at 1500 × g for 10 min. Two pellets (bottles) are pooled, washed with saline (1500 × g, 10 min), and resuspended in 40-mL K2Cr2O7. Secondary gradients are prepared and processed as described earlier, except that the sample overlaid is the 40-mL primary gradient-purified oocyst fraction. Paired pellets collected from the secondary gradients are resuspended to 40-mL K2Cr2O7 and stored at 4°C. If oocysts are desired that have not been exposed to oxidizers such as phenol and K2Cr2O7, simply replace the phenol in Sheather’s solution with water and use saline or PBS, with or without antibiotics, as the oocyst storage medium. Note: Sheather’s solution and the STP solutions that do not contain phenol have a shorter shelf life, even if stored at 4°C, and should be used promptly to avoid growth of contaminating microorganisms.
4.
Cesium Chloride Gradients
Final oocyst purification is accomplished using a simple, micro-scale cesium chloride (CsCl) gradient technique (Figure 20.3) (Arrowood and Donaldson, 1996). The secondary sucrose gradient-purified oocyst fractions are centrifuged (1500 × g, 10 min) and resuspended in 3-mL saline. Siliconized 1.7mL polypropylene microcentrifuge tubes are filled with 750-µL cesium chloride solution (sp. gr. 1.15 = 21.8-g CsCl, 103.3-mL deionized H2O). Oocyst suspension aliquots (500 µL) are carefully layered over the CsCl solution and centrifuged at 16,000 × g (approximately 13,200 rpm) for 3 min in a standard microcentrifuge. The upper 1-mL layers are collected, transferred to clean, siliconized 1.7-mL microcentrifuge tubes, diluted to 1.5 mL with saline, and centrifuged (16,000 × g, 3 min). Pellets from six tubes, corresponding to the original secondary oocyst fraction, are pooled, washed a second time with saline, resuspended in 1-mL 2.5% K2Cr2O7 or antibiotic supplemented saline or PBS, and stored at 4°C. At this point, oocysts are essentially free of gross debris and contaminating microorganisms. The few remaining bacteria are usually nonviable carcasses. Figure 20.1 summarizes the oocyst purification process.
5.
Alternate Methods
A variety of alternative oocyst purification methods are available and preferred by different laboratories or for particular purposes. Conventional and modified salt or sucrose flotation techniques, Percoll® or Ficoll® gradients, cesium chloride gradients, and cellulose filtration methods have been advocated (Riggs and Perryman, 1987; Taghi-Kilani and Sekla, 1987; Rosales et al., 1994; Meloni and Thompson, 1996; Upton, 1997).
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510 D.
Cryptosporidium and Cryptosporidiosis, Second Edition Excystation of Sporozoites
Successful infection of cell cultures requires excystation of sporozoites from oocysts. In its simplest form, this can be accomplished by inoculating purified, intact oocysts into host cell cultures because excystation will proceed as oocysts respond to temperature and chemical triggers naturally occurring in culture media. Traditionally, sporozoite excystation was initiated by suspending oocysts in a trypsin and sodium taurocholate solution (0.25 and 0.75%, respectively) prepared in a buffered salt solution (e.g., PBS or Hanks Balanced Salts Solution [HBSS]), sometimes simultaneously or in ordered sequence. To avoid enzymatic damage to sporozoite surface proteins, 0.75% synthetic sodium taurocholate can be used alone in sterile PBS, HBSS, or a basal culture medium (e.g., DMEM), incubating at 37°C for 10 to 15 min (as a “trigger” before inoculating oocysts into cell culture) or for 45 to 90 min for total sporozoite release before inoculation (verify excystation microscopically). Sporozoites should exhibit “flexing” and may display motility if contact is made with slide, coverslip, or host cell surfaces. If sporozoites are rounded or “stumpy”, or if they disappear or rupture soon after excystation, the excystation medium may be contaminated with residual K2Cr2O7, bleach, or other salts, or may have osmolarity problems. Some investigators treat purified oocysts with sodium hypochlorite (dilute commercial bleach) at a 1:10 dilution in water (or even undiluted bleach!) for approximately 10 min at 4°C or at room temperature. The bleach is washed out (or neutralized with 0.1% sodium thiosulfate), and oocysts are suspended in excystation solution or directly inoculated onto host cells. Bleach treatment “triggers” excystation (or possibly enhances overall excystation rates) and can provide additional “sanitizing” of the oocyst sample to prevent microbial contamination of host cells. If oocysts have been processed using sucrose gradients (with phenol) and stored in K2Cr2O7, microbial contamination will be unlikely. Of course, contaminantfree oocysts can always be “recontaminated” by poor aseptic technique. Unless oocyst stocks are very old (> 6 months) or microbial contaminants are a problem, the bleach step is unnecessary and should be avoided if performing certain disinfection and survival studies as the bleach treatment may damage the disinfectant-weakened or stressed oocysts.
E.
Sporozoite Purification
A variety of methods are available to purify sporozoites: Percoll® gradient separation (Figure 20.3) (Arrowood and Sterling, 1987; Nesterenko and Upton, 1996; Upton, 1997), cellulose ion exchange column chromatography (Riggs and Perryman, 1987; Nesterenko and Upton, 1996; Upton, 1997), or by filtration through 2- or 3-µm polycarbonate track etch membranes (Current, 1990; Gut et al., 1991; You et al., 1998). A caveat regarding purifying sporozoites: they are somewhat fragile and perform best in culture when handled least. Furthermore, excystation in advance of purification is best accomplished with synthetic sodium taurocholate, as crude taurocholate preparations quickly cause sporozoites to “round up” or appear “stumpy” and presumably interferes with invasion potential. The duration of excystation and sporozoite purification should be as short as possible to avoid exposure to oxygen and formation of oxygen radicals in sporozoites because it is likely to negatively impact the invasive and developmental potential of the parasite (Upton, 1997).
1.
Sieving Technique
As mentioned earlier, a simple sporozoite isolation technique is to “sieve” the excystation mixture through polycarbonate track-etch membranes by attaching a syringe to a commercial disposable membrane unit or to a membrane holder loaded with a 2- to 3-µm porosity membrane (Current, 1990; Gut et al., 1991; Deng and Cliver, 1998; You et al., 1998). This style of membrane has discrete pores that sporozoites can move through while intact oocysts and oocyst shells are retained. A gentle, pulsing pressure works well, and once the entire volume has passed through the assembly, the sample can be followed with plain buffer as a flushing step. Depending on the load, the membrane may become blocked and the syringe may be switched to a new holder and membrane assembly until the sample is completely filtered. Between 107 and 108 oocysts and sporozoites may be accommodated by a 25-mm diameter membrane.
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DE-52 Anion Exchange Chromatography
Sporozoites excysted from bleach-treated oocysts (see earlier text) are suitable for purification via cellulose chromatography (bleach-treated oocysts adhere to the matrix, whereas sporozoites pass through) (Riggs and Perryman, 1987; Upton, 1997). Success with this method is variable and may require considerable “fine-tuning” to reduce intact oocyst and oocyst shell contamination.
Percoll® Gradients
3. ®
Percoll has been used extensively for sporozoite and merozoite purification (Arrowood and Sterling, 1987; Taghi-Kilani and Sekla, 1987; Regan et al., 1991; Nesterenko and Upton, 1996). These methods work best with oocysts that have high excystation rates (> 80–90%). The following briefly describes a rapid and convenient method for isolating sporozoites (Nesterenko and Upton, 1996; Upton, 1997). Excyst sporozoites from purified oocysts using a suitable method (see above, but usually performed in a siliconized 1.5-mL microcentrifuge tube). Centrifuge the suspension (5000 × g, 3 min), aspirate the supernatant, and resuspend the pellet in 1.2- to 1.5-mL isotonic Percoll® solution (sp. gr. 1.04) prepared by mixing commercial Percoll® solution into 0.15-M aqueous NaCl at 23% (v/v). Centrifuge the suspension at 5000 × g for 3 min, discard oocyst shells (very top layer of the Percoll® column), transfer the “cloudy” middle layer above the pellet to new microcentrifuge tubes, dilute with ≥ 4 volumes PBS, and centrifuge (5000 × g, 3 min) to pellet sporozoites. Wash sporozoite pellet by resuspending in PBS or sterile culture media and centrifuging (5000 × g, 3 min).
4.
Sporozoite Viability and Infectivity
Estimating sporozoite viability, and by extension infectivity, is advocated using vital dyes (e.g., fluorogenic dyes such as DAPI) (Campbell et al., 1992, 1994; Robertson et al., 1992; Fricker 1994). However, such methods do not always correlate well with infectivity (Korich et al., 1990; Black et al., 1996). It is also clear that excystation and the release of motile sporozoites can be deceptive because treated sporozoites can invade host cells but fail to replicate (Upton and Tilley, 1995; Shin et al., 2001). Nevertheless, excystation provides a convenient measure of oocyst and sporozoite health. If excystation yields morphologically characteristic sporozoites at rates > 60 to 70%, the preparation is likely suitable for culture.
IV.
In Vitro Culture
Figure 20.2 summarizes the process of establishing cryptosporidial infections in cell cultures using cryptosporidial oocysts (or purified sporozoites). In vitro culture is established by inoculating (1) whole oocysts, (2) mixtures of intact oocysts, excysted oocysts, and free sporozoites, or (3) purified sporozoites. The recommended procedure described in the following text is optimized for intact oocysts; substitute sporozoite numbers if using excysted or purified sporozoites as the inoculum. Figure 20.3 illustrates the appearance of life-cycle stages in various microscopic assays.
A. 1.
Oocyst and Sporozoite Preparation Oocyst Age
Oocyst age certainly affects in vitro infectivity, but isolate characteristics, storage conditions, and purification techniques all contribute to the useable “shelf life” of oocysts. Oocysts “pampered” during collection and purification will be useful far longer than oocysts exposed to fluctuating temperatures, overgrowth of contaminating microorganisms, and harsh chemical treatments. A general recommendation is to use oocysts less than 2 months old (after collection from the host). However, oocysts are often useful for as long as 4 to 6 months or more after collection. In vitro cultivation studies should always include age-matched oocyst controls to account for variability in infection efficiency.
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Gradient purified oocysts Oocyst labeling to track in culture Wash oocysts (centrifuge) to remove K2Cr2O7 or antibiotic mixture.
Suspend oocysts in acetate buffer with periodic acid.
Inoculate oocysts directly onto cell cultures or initiate excystation with exposure to culture medium supplemented with synthetic sodium taurocholate and then inoculate onto cell cultures.
Remove periodic acid via centrifugation and suspend oxidized oocysts in acetate buffer with FTSC, incubate 1 hr. Remove unbound FTSC via centrifugation.
Leave inoculum on cells until growth assayed or remove via washing with PBS after approximately 3 hr incubation.
Wash cultures with PBS after 48 to 72 hr (maximum parasite loads) and fix with Bouin’s solution 1 hour. [Replace Bouin's solution with alternative fixative, depending on assay requirements.]
Decolorize (permeabilize) culture monolayer with 70% ethanol. [Skip if using fixative other than Bouin’s solution.]
Replace ethanol with PBS. Block with PBS/BSA or FBS-containing culture medium.
Perform immunoassay: immunofluorescence, ELISA, chemiluminescence…
FIGURE 20.2 Flowchart summarizing the in vitro culture of Cryptosporidium and preparation of infected cell monolayers for parasite life-cycle stage analysis and quantification.
2.
Preinoculation Labeling of Oocysts and Debris with Fluorescent Dye
A very simple fluorescent dye labeling technique facilitates differentiating inoculated oocysts, oocyst walls, and contaminating organic debris (e.g., dead bacterial cells, fungal spores …) from de novo parasite life-cycle stages (Arrowood and Donaldson 1996). To minimize oocyst losses, use siliconized microcentrifuge tubes throughout the method. Briefly, sucrose and cesium chloride gradient-purified oocysts are washed with acetate buffer (0.1-M acetate, pH 5.5, supplemented with 0.85% NaCl) using a microcentrifuge (16,000 × g, 3 min), suspended in acetate buffer supplemented with sodium periodate (10-mM, prepared fresh) for 20 min on ice, microcentrifuge washed (3 min as earlier) with PBS/BSA (0.1-M PBS supplemented with 0.1% BSA), and suspended in 1-mL acetate buffer supplemented with 150 μg/mL fluorescein thiosemicarbazide (FTSC #46985, Sigma Chemical Company, St. Louis, MO; 10× concentrate prepared fresh in DMSO) for 1 h at room temperature, protected from light. Unbound FTSC is removed by washing twice in PBS/BSA using the microcentrifuge. FTSC is bound to the periodic acid-oxidized carbohydrates, yielding highly fluorescent oocyst walls, bacteria, and other carbohydrate-rich organic debris. Sporozoites within oocysts remain unlabeled. By selecting appropriate detecting reagents/fluorochromes, the labeled inocula are easily distinguished from developing parasites using fluorescence microscopy (Figure 20.3). Note: For laboratories that routinely use bleach treatment to trigger or enhance excystation, bleach treatment cannot follow FTSC labeling, as it will destroy the fluorochrome dye. Caution should be exercised if used prior to FTSC labeling as some oocysts will be permeabilized by the bleach treatment, and FTSC will penetrate and label such oocysts internally (including sporozoites and other oocyst contents).
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FIGURE 20.3 (A color version of this figure follows page 242.) Panels A, B, and C illustrate the progressive gradient techniques used to purify C. parvum oocysts. Panel A shows primary (left) and secondary (right) discontinuous sucrose gradients after centrifugation with the light debris (d) at the interface between the sample and the STP 1:4 layer, the oocysts (o) at the interface between the STP 1:4 and STP 1:2 layers, and the pellet (p) of dense debris. Panel B shows examples of the cesium chloride step gradients: the first tube (left) shows the sucrose gradient-purified oocyst sample (s) layered over the cesium chloride column before centrifugation; the second tube (middle) shows the same tube after centrifugation with the oocysts (o) at the interface between the sample and the cesium chloride and a small pellet (p) of dense debris at the bottom; the third tube (right) shows the purified pellet of oocysts (o) after the saline wash step. Panel C shows an example of a Percoll® gradient used to isolate sporozoites (sp) and empty or partially empty oocyst walls (w). Panel D shows a differential interference contrast (DIC) view of representative C. parvum oocysts purified using the sucrose and cesium chloride gradient techniques. Panel E shows purified sporozoites labeled with a monoclonal antibody specific for a sporozoite surface protein (C3B4, reacts with the 23-kDa surface antigen); this view includes a secondary antibody conjugated with fluorescein (green). Panel F shows the same antibody labeling of merozoites emerging from a meront in a paraffin section of infected mouse ileum. Panels G and H show, respectively, the DIC view and immunofluorescent view (C3B4 labeling) of budding merozoites in a cell culture preparation. Panels I, J, and K show, in sequence, the DIC view of a late trophozoite/early meront, DAPI nuclear staining of the host cell and the parasite nuclei (above host nucleus, see arrow) and, finally, the intracellular label of the parasite by the panreactive monoclonal antibody C3C3 (conjugated with the red fluorescent dye Cy3). Panel L shows C. parvum growth in MDCK cells at 24 h post inoculation (PI) where the oocyst (green) has been labeled directly with FTSC and the developing stages (mostly early trophozoites/Type II meronts) have been labeled with the monoclonal antibody C3CCy3. Panel M shows MDCK cells at 48 h PI; note the obvious FTSC-labeled oocyst and the distinct C3C3-Cy3 labeling of the gamonts and meronts. Scale bar applies to panels D through M.
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Cryptosporidium and Cryptosporidiosis, Second Edition Taurocholate Treatment
Resuspend FTSC-labeled oocysts (or bleach-treated oocysts) in culture medium used with host cells (supplemented with 0.75% synthetic sodium taurocholate), enumerate using a hemacytometer, and inoculate onto cell cultures. If the dilution of the parasite inoculum into the culture medium is ≥ 1/15 (i.e., the final concentration of the taurocholate in the culture medium is < 0.05%), there is no need to wash the taurocholate out of the inoculum. Use of taurocholate is optional but may enhance excystation and subsequent parasite development.
4.
Inoculum Density
Oocysts are generally inoculated at a ratio of 500 to 2000 per mm2 of host cell monolayer (Table 20.2 presents routine densities for a variety of culture containers). Alternatively, they could be inoculated at a ratio of 1:1 or 1:2 (oocysts to host cells) (Rasmussen et al., 1993; Eggleston et al., 1994; Woods et al., 1995; Upton 1997). Inoculum densities much higher than these promote monolayer damage and sloughing. When intact oocysts are inoculated, it is recommended to gently “wash” cells with PBS or culture media 3 h after inoculation and replace with fresh culture media. This step removes free sporozoites, intact oocysts, and oocyst shells that, nevertheless, may adhere to the monolayer and contribute to the “background” signal in growth assays (see the following text). More importantly, oocysts or their contents may inhibit parasite growth by a toxic mechanism against the host cells if they are left in the culture preparations (Eggleston et al., 1994). This phenomenon is clearly apparent with oocysts of coccidia, especially species of Eimeria (Patton, 1965; Fayer and Hammond, 1967; Doran, 1970). Whether this is a significant problem with Cryptosporidium oocysts is less clear. Host cell damage appears to be directly related to successful infection, especially at high parasite loads. Heat-inactivated oocysts inoculated at 10 times the normal inoculum density do not impair MDCK host cell monolayers even if left on the host cells throughout the culture period (data not shown). Oocyst disinfection and survival studies may require leaving oocysts in contact with the monolayer throughout the culture period to verify excystation has not simply been delayed by the treatment. Include appropriate heat-inactivated controls (50°C, 60 min or 100°C, 15 s) at matched or higher densities to account for possible inhibition or toxicity by the inoculum. TABLE 20.2 Recommended Oocyst or Sporozoite Inoculum Densities (by Area) for Commonly Available Culture Chambers and Flasks Container
Unit
Dimensions
Area (cm2)
Media Volume (mL)
1-well slide 2-well slide 4-well slide 8-well slide Leighton tube 24-well plate 6-well plate Costar insert Small flask Large flask
well well well well tube well well well flask flask
2.0 × 4.3 cm 2.1 × 2.1 cm 0.9 × 2.1 cm 0.9 × 0.9 cm 0.9 × 5.5 cm 1.5 cm dia. 3.6 cm dia. 2.4 cm dia. 5 × 5 cm 7.5 × 10 cm
8.6 4.4 1.9 0.8 5.0 1.8 10.2 4.5 25 75
4–5 1.0–1.5 0.4–0.5 0.3–0.4 2–3 1–2 3–4 1.0–1.5 5–6 15–20
a
Sporozoites/ Oocystsa 1 × 106 5 × 105 2 × 105 1 × 105 5 × 105 2 × 105 1 × 106 5 × 105 2.5 × 106 7.5 × 106
A range of 500 to 2000 oocysts per mm2 is typical, depending on the age of the oocysts. Very fresh oocysts should be inoculated at the lower end of the range, whereas aged oocysts (> 4 months) can often be inoculated at the upper end of the range. Exceeding these oocyst or sporozoite densities may result in monolayer damage and sloughing from culture surfaces. An alternative density calculation can be made by inoculating oocysts at a 1:1 or 1:2 ratio relative to host cells in the monolayer.
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Efficiency of infection is typically 5 to 10%, depending on the number of oocysts (or sporozoites) inoculated, the excystation rate, the typical yield of sporozoites, and the common observation that not all sporozoites will attach and invade host cells (some attach and detach without invading) and some sporozoites invade but fail to replicate. Parasite numbers reach a maximum approximately 48 to 72 h after inoculation. A general rule of thumb is that developing stage numbers will equal (or roughly double) the number of original oocysts/sporozoites in the inoculum. Some investigators observe increased infection efficiencies if the original inocula are gently centrifuged onto the host cell monolayers (Weir et al., 2001). The distribution of developing parasites across host cell monolayers is dependent on adequate mixing of the inoculum into the culture medium overlaying the host cells at the time of inoculation. Similar to coccidia, Cryptosporidium will often be distributed in “patchy” foci, a consequence of the motility and rapid (nearby) invasion of host cells by merozoites as they are released from Type I and Type II meronts.
B. 1.
Host Cells and Culture Environment Host Cell Selection
Numerous host cell lines have been tested and advocated for cultivating C. parvum (see Table 20.1). Human ileocaecal adenocarcinoma (HCT-8), MDCK, and human colonic adenocarcinoma (Caco-2) cells are the most commonly used cell lines. HCT-8 and MDCK cell lines are particularly useful as they are easy to maintain as stock cultures, form consistently confluent monolayers, are contact inhibited (growth slows as the monolayer becomes confluent), and are highly adherent to culture surfaces, including most plastic and glass cultureware. The latter feature is valuable for microscopic studies and for assays dependant on monolayers remaining in place throughout culture and parasite detection methodologies. Both cell lines require routine trypsinization (trypsin 0.5 mg/mL, EDTA 0.2 mg/mL in HBSS) to be released from culture surfaces and to form disperse (mostly single-cell) suspensions before subculture. Uniform, synchronized, differentiated cells are ideal targets for sporozoites, and flat monolayers facilitate microscopic assays. Transferring clumped cells during subculture results in “piled” or “mounded” cell islands with uneven (3-dimensional) parasite distribution. Initiate subculture of HCT-8 or MDCK cells by splitting and inoculating cells into selected culture chambers at ratios that will yield confluent monolayers within 48 to 96 h. If Caco-2 cells are used, subculture is initiated similar to the HCT-8 and MDCK cells, but once cells become confluent, growth medium can be replaced by maintenance medium (lower concentration of FBS) so that cells can be maintained for extended periods (>7 days) to fully differentiate (Favennec et al., 1994).
2.
Atmosphere Selection
A variety of gaseous environments have been tested to support cryptosporidial growth (Upton et al., 1994b). A simple 5% CO2/95% air mixture incubator at 37°C conveniently supports C. parvum growth in HCT-8, MDCK, and Caco-2 cells.
3.
Culture Media
As described earlier and in Table 20.1, a variety of culture media and supplements have been studied to find optimal conditions that support C. parvum growth in various cell lines. The customized RPMI 1640-based medium described by Upton et al. (1995) yielded optimal parasite growth in HCT-8 cells compared to medium with standard supplements. A simple alternative is Ultraculture® (Cambrex Bio Science, Walkersville, MD), a synthetic medium that supports comparable or better parasite growth in both HCT-8 and MDCK cells (Figure 20.4). The medium, as supplied, has a long shelf life and requires only the addition of L-glutamine before use. No supplementation with FBS or other components is necessary.
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24 h PI 48 h PI Meronts and Gamonts/mm
2
2500
2000
1500
1000
500
0 MDCK Ultra
HCT-8 2 Ultra
HCT-8 RPMI
2 d a ys
MDCK Ultra
HCT-8 4 Ultra
HCT-8 RPMI
MDCK Ultra
4 d a ys
HCT-8 8 Ultra
HCT-8 RPMI
8 d a ys
Host cell age (before inoculation) FIGURE 20.4 Cryptosporidium parvum parasite production in MDCK and HCT-8 cells using commercially available Ultraculture® medium (Arrowood et al., 1994) compared to customized RPMI medium (Upton et al., 1995); the effect of host cell age at the time of inoculation (days after splitting and seeding into culture chambers) is also shown. Data are presented for 24 and 48 h post inoculation (PI). Parasite production generally peaks between 48 and 72 h PI.
C.
Quantification of Developing Parasites in Culture
In vitro development of C. parvum can be assessed using a variety of tools, including microscopy, immunoassays, and various molecular biological methods targeting parasite nucleic acids.
1.
Microscopic Enumeration
Microscopic assays have been widely used in cryptosporidial culture studies. Host cells grown on coverslips or in culture chambers with microscope slides or coverslip bases (e.g., LabTek® Chamber Slides® or Chambered Coverglass®, Thermo Fisher Scientific, Inc., Waltham, MA) are inoculated, incubated, fixed using methanol, acetone, aldehydes (formalin, paraformaldehyde, or glutaraldehyde), or Bouin’s solution, and stained with Giemsa, hematoxylin and eosin, Hoechst 33258, Papanicolaou, Alcian blue, Ziehl–Neelsen or other acid-fast dyes (Naciri et al., 1986; Arrowood, 1988; McDonald et al., 1990; Flanigan et al., 1991; Gut et al., 1991; Kuhls et al., 1991; Marshall and Flanigan, 1992; Martinez et al., 1992; Doyle et al., 1993; Rasmussen et al., 1993; Rosales et al., 1993; Wiest et al., 1993; Rosales et al., 1994; Deng and Cliver, 1998). Parasites are quantified as a ratio of parasites to host cells, numbers per field or unit area, or using variations on most probable number (MPN) techniques (e.g., number of fields positive across a given area rather than counting individual parasites). Stained or labeled slides provide a permanent or long-term record of results and can be repeatedly evaluated or examined over a flexible time frame. Nevertheless, direct observation of cultures or stained or labeled parasites can be confounded by difficulty in distinguishing parasites stages from host cells, distinguishing parasite stages from one another (e.g., inoculum versus de novo stages) or losses of host cells and parasites during assay processing. Use of parasite-specific antibodies in bright-field immunostaining or immunofluorescent labeling of parasite stages provides a valuable alternative to less-specific chemical stains (Arrowood et al., 1994;
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Arrowood et al., 1996; Slifko et al., 1997; Slifko et al., 1999). MDCK cells are preferred for immunofluorescence in coverglass-bottomed culture chambers (LabTek® Chambered Coverglass®) because cells grow flat and the developing parasites are found within a narrow focal plane, facilitating image capture for subsequent software-assisted analysis. By comparison, HCT-8 cells grow much more 3-dimensionally (“hills and valleys”), complicating image capture-based microscopic analyses at high magnification. However, both cell lines are well suited for low- and medium-power magnification (200 to 400×) examination, real-time microscopy, and for nonmicroscopic assays. LabTek® Chambered Coverglass® chambers facilitate microscopic observation of live cells (up to 1000× total magnification) and after fixation and labeling can be scanned directly on an inverted microscope, or the chambers can be inverted onto microscope slides for observation on a standard “upright” microscope. A variety of antibody reagents (both polyclonal and monoclonal) have been used for immunofluorescent visualization of life-cycle stages (Arrowood, 1988; Arrowood et al., 1989; Arrowood et al., 1991; Tilley et al., 1991; Arrowood et al., 1994; Favennec et al., 1994; Griffiths et al., 1994; Gut and Nelson, 1994; Arrowood et al., 1996; Rochelle et al., 1997; Slifko et al., 1997; Deng and Cliver, 1998). Figure 20.3 illustrates the value of immunofluorescent labeling in visualizing developing stages in culture: some parasites are quite striking in appearance, but not all parasite stages are easily identified by DIC or bright-field imaging. Use of immunofluorescent labeling makes identification much easier, but if the antibody does not react with all developing stages (note in Figure 20.3, panels G and H, that only the surface of the budding merozoites are labeled, whereas the rest of the meront is not), some stages can be missed when evaluating cultures. If a pan-reactive monoclonal or polyclonal antibody is used, especially if combined with the FTSC labeling of the inocula, all de novo stages can be visualized and enumerated. The following technique is straightforward and yields intensely labeled parasites suitable for direct imaging and image capture-based studies. Immunofluorescence labeling of parasite cultures is initiated by rinsing inoculated cultures with PBS, fixing with Bouin’s solution for 1 h and decolorizing with 70% ethanol (five changes) for 1 h. Bouin’s fixation enhances monolayer adhesion to culture chamber surfaces, especially glass. Monolayers are subsequently washed with PBS, labeled for 60 min at room temperature (RT) with neat hybridoma culture supernatant containing a Cryptosporidium-specific monoclonal antibody (e.g., C3C3) or with appropriately diluted antibody directly labeled with a fluorescent dye (e.g., fluorescein or indocyanine [(Cy3®, GE Healthcare)]. If nuclear staining is desired, primary antibody labeling solutions are supplemented with DAPI (0.25 μg/mL, 4, 6-diamidino-2-phenylindole). Cultures with unlabeled primary antibody are subsequently labeled (60 min at RT) with an appropriate secondary reagent (e.g., fluorescein or Cy3-labeled goat antimouse immunoglobulin). Labeled cell cultures are washed with PBS/BSA, coverslipped with mounting medium (supplemented with an antiquenching agent), and observed with a UV microscope. A suitable, long-term mounting medium can be prepared using 2.4-g polyvinyl alcohol (Sigma P8136), 6-g glycerol, 6-mL H2O, 12-mL 0.2-M Tris buffer (pH 8.5), and 2.5% (w/v) of the antiquenching agent DABCO (1,4-diazabicyclo-[2.2.2]-octane) (dissolve in the order listed and warm briefly to 50°C to dissolve completely) (Harlow and Lane, 1988). The unused mounting medium can be stored at –20°C for several months. Life-cycle stages are identified by reactivity with monoclonal antibody, by size and morphology, and, if DAPI employed, by distinctive nuclear staining. Only meronts and gamonts (≥ 3 µm) are enumerated to avoid counting nonviable, adherent sporozoites or merozoites. Meronts and gamonts can be quantified as number/mm2 using a variation of the formula reported by Augustine (1980): stages/mm2 = [(total # stages in 30 fields)/(K × PC)], where PC is the average percent confluence for each field and the constant K represents the area in mm2 of 30 microscopic fields (Arrowood et al., 1994,1996). Alternatively, the entire monolayer is scanned at 200× magnification and scored per field as containing or not containing life-cycle stages to calculate the percentage of fields positive. See appropriate publications for detailed descriptions of parasite enumeration techniques (Arrowood et al., 1994, 1996; Heindel et al., 1999; Slifko et al., 1999). An advantage of microscopic methods is the ability to identify very small numbers of developing parasites (even single meronts and gamonts). Trophozoites at an early stage of development are virtually indistinguishable from newly invaded sporozoites or merozoites. Some sporozoites and merozoites never invade host cells, but may attach to cell surfaces and “round-up” and appear to have entered host cells, whereas others invade host cells and fail to develop further. One way
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to avoid counting these nonpropagating forms is to simply not count developing stages smaller than 3 µm in diameter, thus restricting quantification to maturing meronts and gamonts. Nonmicroscopic methods often have a higher threshold of detection or may risk detecting antigens or nucleic acids that are part of the parasite inoculum. Figure 20.3 presents several microscopic views of C. parvum in culture including differential interference contrast view of sporozoites, asexual and sexual stages, and DAPIlabeled parasite nuclei. Differentiating the inoculum (oocysts and oocyst walls) from de novo parasites using the FTSC labeling technique described earlier is illustrated. When combined with contrasting antibody-fluorescent dye labels (e.g., FTSC labeling method described earlier), differentiation of lifecycle stages, including culture-derived oocysts, is readily accomplished.
2.
Enzyme Immunoassays (EIA)
Enzyme immunoassays provide an effective alternative to microscopic methods for evaluating parasite development and can be most valuable in large-scale assays adapted to 96-well culture plates (Upton et al., 1994c; Woods et al., 1995; You et al., 1996a, 1996b). Disadvantages include the lack of a permanent record (that can be reexamined), inability to distinguish developmental stages (primarily a limitation of the detecting antibody reagent), and propensity to detect the part of the inoculum (sporozoites and oocysts) that adheres to the host cell monolayer and is not removed by wash steps or medium changes. A method using HCT-8 cells (Upton, 1997) is described briefly. HCT-8 cells are split into culture wells (96-well plate) and allowed to form confluent monolayers (24 to 96 h). Culture wells are seeded with 2 to 3 × 104 purified oocysts. Depending on the assay, bleach treatment of oocysts (see earlier text) can be incorporated to enhance excystation and reduce contamination. Rinse wells with PBS after incubation for 90 min and replace with fresh growth medium (supplemented as necessary with test compounds). Incubate 48 to 72 h at 37°C in a 5% CO2 incubator. Add 100-µL 4% formalin to each well to stop parasite development and to “fix” the monolayer. Fixed monolayers are washed 3× with PBS, blocked with PBS supplemented with 1% BSA and 0.002% Tween-20, and incubated for 30 to 60 min with anti-Cryptosporidium antisera or monoclonal antibodies diluted in PBS blocking buffer (or equivalent). The primary antibody is removed from the wells, the plate is washed with a buffer, and a secondary reagent conjugated with a reporter (e.g., goat antirat antiserum conjugated to horseradish peroxidase) is added and incubated for 20 to 60 min. Plates are washed again and enzymatic color development completed by adding appropriate substrate (e.g., 3,3,5,5-tetramethylbenzidine). Plates are read using a microplate reader at an appropriate wavelength depending on the enzyme and substrate used.
3.
Molecular Methods and Other Alternatives
Incorporation of exogenous 3H-uracil as a measure of coccidian parasite development has been successful with Toxoplasma and Eimeria (Pfefferkorn and Pfefferkorn, 1977; Schwartzman and Pfefferkorn, 1982; Schmatz et al., 1986). Despite the initial reports of success with Cryptosporidium (Upton et al., 1991; Eggleston et al., 1994), the method has not been adopted as a routine assay by other laboratories. Whether this is an indication that Cryptosporidium inefficiently incorporates the exogenous 3H-uracil or preferentially scavenges host cell uracil is unclear. Molecular biology techniques, including polymerase chain reaction (PCR) assays, have been successfully used in assays of cryptosporidial growth (Rochelle et al., 1996, 1997, 2002; Jenkins et al., 2003; Keegan et al., 2003; Di Giovanni and LeChevallier, 2005). The complexity of molecular targets, probe construction, and variety in assay conditions precludes presenting a simple method description. The reader is directed to the respective publications for assay details. Nevertheless, results from comparisons between animal models and either immunoassay-based cell culture or molecular assay-based cell culture showed strong agreement between the murine model (“gold standard” for evaluating oocyst viability and infectivity) and cell-culture-based assays. Given the great sensitivity of PCR-based assays, detecting adherent but nonviable parasites from the inoculum is a potential problem. Reverse transcriptase PCR to detect parasite mRNA is probably a better assay choice because it should reduce detection of nonreplicating parasites (Rochelle et al., 1997, 2002; Jenkins et al., 2003). Selection of target mRNA is important to avoid mRNA pools in the oocyst and sporozoites of the inoculum.
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TABLE 20.3 Cell Culture Abbreviations, Acronyms, Descriptions, and Definitions Abbreviation
Name/Definition/Use
BFTE BHK BS-C-1 Caco-2 CAM HCT-8 HFL HRT HT29.74 Intestine 407 L929 LGA MDBK MDCK MRC-5 PCK PK-10 RL95-2 T84 THP-1 ATCC BSA CCL CRL DABCO DAPI DMEM DMSO ECACC FBS Ficoll® FTSC HBSS HEPES HTB IU K2Cr2O7 Leighton tube MEM PBS PCR Percoll®
Bovine fallopian tube epithelial cells Baby hamster kidney cells African green monkey kidney cells Human colonic adenocarcinoma cells Chorioallantoic membrane (chicken egg) Human colonic tumor (human ileocecal adenocarcinoma) cells Human fetal lung cells Human rectal tumor cells Human colon adenocarcinoma cells Human embryonic intestine cells Mouse fibroblast cells Lewis rat small intestine carcinoma cells Madin Darby bovine kidney cells Madin Darby canine kidney cells Secondary human lung fibroblast cells Primary chicken kidney cells Porcine kidney cells Human endometrial carcinoma cells Human colon carcinoma cells Human monocyte, nonadherent cells American Type Culture Collection Bovine serum albumin Certified cell line Cell repository line 1,4-diazabicyclo-[2.2.2]-octane, immunofluorescence antiquenching agent 4, 6-diamidino-2-phenylindole, nuclear stain Dulbecco’s modified Eagle’s medium Dimethyl sulfoxide, used in cryopreservation media European Collection of Cell Cultures Fetal bovine serum Neutral, highly branched, hydrophilic polymer of sucrose used in gradient purification methods Fluorescein thiosemicarbazide, oocyst surface-labeling reagent Hank’s balanced salts solution N-2-Hydroxyethylpiperazine-N-2-ethanesulfonic acid, buffer used in cell culture Human tumor bank International units Potassium dichromate (2.5% aqueous working concentration for storage of oocysts at 4°C) Glass culture tube with a flattened side in which a coverglass can be positioned to support cell growth Minimal essential medium Phosphate-buffered saline Polymerase chain reaction, DNA detection assays Polyvinylpyrrolidone (PVP)-coated silica particles (15–30 nm diameter) used in gradient purification methods for cells Roswell Park Memorial Institute medium
RPMI
A recent study described the enumeration of Cryptosporidium-infected cells in in vitro cultures using flow cytometry (Mele et al., 2003). Infected cells were detected using either a parasite-specific monoclonal antibody recognizing the “SA35” antigen or the Vicia villosa lectin (VVL labeled with biotin), which binds to parasite glycoproteins in the parasitophorous vacuole. The primary antibody (or lectin) was labeled with a fluorescent secondary reagent and evaluated on a flow cytometer. The authors reported narrower coefficient of variance for the flow cytometry results compared to conventional microscopy using the fluorescent reagents.
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Cryptosporidium and Cryptosporidiosis, Second Edition Culture in Avian Embryos
Joining the ranks of coccidia successfully grown in avian embryos (Doran, 1973), C. parvum and C. baileyi have been grown in avian embryos (primarily chicken eggs) (Current and Long, 1983; Naciri et al., 1986; Lindsay et al., 1988). The report describing C. muris growth in avian embryos apparently incorrectly identified C. parvum as C. muris (Naciri et al., 1986). Anecdotal accounts of C. parvum cultivation in avian embryos suggest that the source of eggs may be a significant factor in the success of parasite growth. The numerous cell culture models for growing C. parvum and C. hominis have essentially obviated the need for avian embryos in laboratory studies. However, avian embryo culture of C. baileyi is still valuable given the lack of in vitro cell culture models for this species (Lindsay et al., 1988). Briefly, 10- to 12-day-old avian embryos are candled with a bright light to identify areas for parasite injection away from major blood vessels. The target injection site on the shell is marked, sanitized by swabbing with 70% ethanol, and punctured with an 18- or 20-gauge needle. If desired, antibiotics (100IU penicillin and 1- to 10-mg streptomycin) can be injected 24 h prior to parasite injection to reduce the risk of microbial contaminant overgrowth. Approximately 1 to 4 × 105 excysted sporozoites or intact oocysts are injected into the allantoic cavity, the puncture site sealed with hot wax or tape, and the egg incubated at 37 to 38°C (C. parvum) or 40 to 41°C (C. baileyi) for 6 to 7 days. After the incubation period, the egg is opened, and the chorioallantoic membrane (CAM) is removed for direct examination, parasite harvesting (primarily oocysts), or fixed and processed for routine tissue sectioning and staining (Lindsay et al., 1988). Oocysts recovered from the CAM are suitable for a variety of laboratory studies.
V.
Concluding Comments
Great strides have been made in producing in vitro culture models for Cryptosporidium parasites that are useful for assessing potential drug therapies, oocyst disinfection methods, gene expression, and a variety of biological characteristics. Nevertheless, more research is needed to establish in vitro culture conditions that support the efficient production of viable and infectious oocysts and methods for continuous parasite development (which would facilitate the establishment of clonal parasite stocks). A continuing, urgent need is for cryopreservation methods effective for oocysts, sporozoites, or developing stages of Cryptosporidium.
References Adams, R.B., Guerrant, R.L., Zu, S.X., Fang, G.D., and Roche, J.K. 1994. Cryptosporidium parvum infection of intestinal epithelium: morphologic and functional studies in an in vitro model. J. Infect. Dis. 169, 170–177. Akiyoshi, D.E., Feng, X., Buckholt, M.A., Widmer, G., and Tzipori, S. 2002. Genetic analysis of a Cryptosporidium parvum human genotype 1 isolate passaged through different host species. Infect. Immun. 70, 5670–5675. Angus, K.W., Sherwood, D., Hutchison, G., and Campbell, I. 1982. Evaluation of the effect of two aldehydebased disinfectants on the infectivity of faecal cryptosporidia for mice. Res. Vet. Sci. 33, 379–381. Arrowood, M.J. 1988. Cryptosporidium: oocyst production and hybridoma generation for examining colostrum and monoclonal antibody roles in cryptosporidial infections. University of Arizona. Arrowood, M.J. and Donaldson, K. 1996. Improved purification methods for calf-derived Cryptosporidium parvum oocysts using discontinuous sucrose and cesium chloride gradients. J. Eukaryot. Microbiol. 43, S89. Arrowood, M.J., Mead, J.R., Mahrt, J.L., and Sterling, C.R. 1989. Effects of immune colostrum and orally administered antisporozoite monoclonal antibodies on the outcome of Cryptosporidium parvum infections in neonatal mice. Infect. Immun. 57, 2283–2288.
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You, X., Arrowood, M.J., Lejkowski, M., Xie, L., Schinazi, R.F., and Mead, J.R. 1996a. A chemiluminescence immunoassay for evaluation of Cryptosporidium parvum growth in vitro. FEMS Microbiol. Lett. 136, 251–256. You, X., Schinazi, R.F., Arrowood, M.J., Lejkowski, M., Juodawlkis, A.S., and Mead, J.R. 1998. In-vitro activities of paromomycin and lasalocid evaluated in combination against Cryptosporidium parvum. J. Antimicrob. Chemother. 41, 293–296. You, X.D., Arrowood, M.J., Lejkowski, M., Xie, L.T., Schinazi, R.F., and Mead, J.R. 1996b. In vitro evaluation of anticryptosporidial agents using MDCK cell culture and chemiluminescence immunoassay. J. Eukaryot. Microbiol. 43, 87S.
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Index
A Acid flocculation, oocyst, 183 Actin avian Cryptosporidium species, 411 avian genotypes, 412 C. baileyi, 407 C. galli, 410 biochemistry, 66 drug targets, 70 genotyping tools, 122 PCR primers, 121 pigs, 471 typing and subtyping methods, 202 Actin-binding proteins, 241 Acute infections, 1 Adaptation to host, see Host, adaptation to Adaptive immune response, see T-cell mediated immunity Aerobic digestion, sludge, 378 Age, host case surveillance, 339 cattle, 452 beef, 457 clinical features and pathology, 453 dairy, 379, 456, 459–460 and prevalence, 455, 457, 461, 462 and species and genotypes infecting, 457, 459–460 and species shed by, 379 clinical disease and pathology, 236, 237–239 children, 237–238 elderly, 238–239 dogs, 441 epidemiology descriptive and analytical investigations, 97 and gender, 86, 87 previous infection and age of children as factor in antibody levels, 86 and susceptibility, 86 horses, 444 molecular epidemiology animal cryptosporidiosis, 132–133 endemic human cryptosporidiosis, 139–140 human infections, 135 role of animals in transmission to humans, 138 pigs, 468, 469–470 sheep, 462 Age, oocyst, 511 Agricultural animals, see Livestock/agricultural animals
Agricultural practices, and endemic human cryptosporidiosis, 139–140 Agricultural products, foodborne disease sources, 297 Albumin, bovine serum, 195 Algal genes, in nuclear genome, 46 Alinia®, see Nitazoxanide (Alinia®) α/β receptor positive T-cells, 214 Amino acid metabolism, 60, 65–66 Amino acids, 259 Amino acid transporters, 46 Ammonia disinfectants, 32, 34, 399 Amphibians, 10, 11, 388–389 Amylopectin, 28 Anaerobic digestion dairy cattle manure, 379–380 sludge, 378 swine manure, 382 Anaerobic metabolism, 46 Analytical investigations epidemiology, 97 molecular epidemiology, 84 Animal cryptosporidiosis, see also Cattle; Farm animals; Livestock/agricultural animals; Pig(s); Sheep chemotherapy and prophylaxis, 265–279 miscellaneous, 276 rodents, 266–273 ruminants, 274–275 in vitro, 277–278 diagnostic methods, 201 epidemiology, 83 age and susceptibility, 86 C. hominis infections, 87 molecular epidemiology, 130–133 species and genotypes in mammals, birds, reptiles, and fish, 130–132 transmission routes, 132–133 wild animals, see Wild animals zoo animals, 426–427 Animal models, 485–493 C. parvum and C. hominis, 145–146 and host range, 493 parasite propagation and oocyst production, 486–487 calves, 507 mice, 506–507 for research, 487–492 agents for treatment and prevention, 28, 31, 488–491 versus cell culture, 492 genetics of cryptosporidia, 491–492
527
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water treatment testing, 492 Animal-specific genotypes, epidemiology, 83 Animal transmission/zoonotic disease, 120 epidemiology risk factors, 87 sources, 88 sporadic disease, 97 transmission routes, 93 transmission routes, C. parvum, 89 molecular epidemiology, 150 C. parvum phylogenetic relationships, 124 C. parvum population genetics, 129 endemic human cryptosporidiosis, 140 human infections, 134 population genetics, significance of, 130 role of animals in transmission to humans, 138–139 source tracking, 146–147 transmission routes, 120, 138–139 transmission routes, 3, 26 wild mammals as reservoir, 428 Animal waste dairy, 379 recreational water contamination, 357 Anion exchange chromatography, sporozoite purification, 511 Annotation, 45, 46–47, 59, 60, 61 Anthroponotic disease, see Human (anthroponotic) transmission Anti[retro]viral therapy, 151, 263, 264; see also HAART therapy molecular epidemiology, 150 in vitro studies, 278 Antibodies birds, 397–398 cats, domestic, wild, and feral, 421–422 cattle, 454 diagnostic methods, 191–193 commercially available, 185 enzyme-linked immunoelectrotransfer blot (EITB), 191–192 enzyme-linked immunosorbent assay, 192–193 immunomagnetic separation (IMS), 184 epidemiology asymptomatic infection, 81 human volunteer studies, 98 previous infection and age of children as factor in, 86 seroepidemiology/population immunity, 85–86 immune response, 225 from immunized chicken, 224 maternal, 86 parasite-specific, 222–224 immunization-induced, 223–225 infection-induced, 222–223 polyclonal immunization with crude antigen, 223–224 immunization with recombinant antigen, 224 recreational water use and, 341 reptiles, 391
Antigen capture methods, 190 Antigens diagnostic methods, 188–190 enzyme-linked antibodies, 189–190 fluorescence-labeled antibodies, 188–189 oocyst negative stool samples, 183 extracellular, 46 membrane proteins, 67–68 polyclonal antibodies from immunization with crude antigen, 223–224 recombinant antigen, 224 seroepidemiology, 85 surface proteins, 49 T-cell priming and T-cell memory, 216–217 Antimicrobial peptides, 211 Apicomplexa biochemical divergence from, 58 biochemistry apicoplast, 63 biochemical divergence of Cryptosporidia from, 57–58 carbohydrate and energy metabolism, 61 DNA replication, 69 fatty acid metabolism, 62, 63, 64 highly streamlined metabolism, 58, 59 membrane proteins and transporters, 68 nucleotide metabolism, 62 polyamine metabolism, 65 structural proteins, 66 coding capacity, 46 comparative genomics, 48–49, 51–52 expressed sequence tags, 44, 45, 46 functional genomic analysis, 47, 48 genome properties, 45–46 genomic sequences, 45 intracellular and horizontal gene transfer, 51, 52 life cycle, 4, 6 excystation, 13, 47 sporozoites, 22 PCR nonspecific amplification, 122 PCR primers, 121 phylogenetic analysis, 49–51 phylogenetic relationships, see Evolutionary/phylogenetic relationships taxonomy, 4 Apicoplast, absence of, 45 Apoptosis, epithelial cells, 242 AP-PCR, 122 Arbitrarily primed polymerase chain reaction (AP-PCR), 122 Architectures, domain, 49 Arginine decarboxylase, plant-type, 65 Arginine-supplemented diet, mice, 221 Arlequins (population genetics program), 169 Artiodactyla, 5 AseI, 199, 200 Asexual multiplication (merogony), 16, 18, 19, 20 Asymptomatic/subclinical infection and case surveillance, 339 C. parvum versus C. hominis, 146 epidemiology, 81
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Index horses, 444 infectivity trials, 243 molecular epidemiology, 146, 150 oocyte shedding, detection limits, 178 recreational water contamination, 337 in reptiles (snakes), 389 Atovaquone (Mepron®), 263 ATPases, 46, 68 ATP-binding cassette transporters, 46 Attachment for feeder organelle, 14, 15, 23–24 Attenuation serial passage and virulence, 454 for vaccines, gamma irradiation of oocysts, 225 AUCP-1 strain, 453–454 Auramine phenol, 186–187 comparison of diagnostic methods, 191 oocyst staining methods, 186–187 Avian cryptosporidia, see also Birds evolutionary relationships of avian isolates, 404 human infections, 134 Avian embryo culture, 520 C. baileyi, 398–399 chorioallantoic membrane culture, see Chorioallantoic membrane (CAM) culture Avian genotypes, 410–413 duck genotype, 412 Eurasian woodcock, 412 evolutionary relationships of avian isolates, 404 genetic similarities at SSU rRNA locus, 407 goose genotypes, 412–413 oocyte morphometry, 400 types I-IV, 410–412 Azidothymidine, 260 Azithromycin, 257, 258, 259, 260–261 mice, 267, 268 rats, 270, 272 in vitro studies, 277
B Baboons, 425 Bacterial genes horizontal (interspecies) gene transfer, 49, 51 in nuclear genome, 46 Bacterial-type enzymes, drug targets, 70 Bactrian camels, 5, 392, 426, 429, 471 Bandicoots, 5, 424 Barium, diagnosis in reptiles, 390 Bats, 424 Bear genotype, 8 evolutionary relationships of avian isolates, 404 Beef cattle, 452 manure, waste management, 381 prevalence, nonmolecular methods, 457, 458 species and genotypes, molecular methods, 460, 461 Benzeneacetonitrile derivatives, 261–262 β-defensin, 211 β-tubulin, 69
529 drug targets, 70 typing and subtyping methods, 202 Beverages, oocyst detection, 295 Biaxin®, Biaxin XL® (clarithromycin), see Clarithromycin Bilbies, 427, 428 Bile, passive protection in rodents, 222–223 Biliary system involvement, 239–240, 241 azithromycin tissue concentrations, 257 immunization and, 224 in immunocompromised host, 256 pathophysiology, 242 pigs, 467–468 Biochemistry, 4, 57–71 amino acid metabolism, 65–66 DNA and RNA metabolism, 58–59 energy and carbohydrate metabolism, 59–61 fatty acid metabolism, 63–65 genomics absence of nucleotide synthetic pathways, 46 databases, 48 functional genomic analysis, 47 nucleotides, as phylogenetic mosaic, 52 nucleotide salvage pathways, absence of biosynthetic pathways, 51 highly streamlined metabolism, 58–59 membrane proteins and transporters, 67–68 nucleotide metabolism, 62 polyamine metabolism, 65 potential drug targets, 69–70 species identification, 7 structural proteins, 66–67 technical perspectives related to biochemical studies, 70–71 Biodrying, dairy cattle manure, 380 Bioinformatics databases, 52–53 “-omics” resources, 47, 48 Biological treatment, waste management, 376 Biology, 1–35 C. baileyi, 403, 404 C. galli, 407–409 C. meleagridis, 399–400 history, 2–4 host specificity, 7–10 life cycle, 11–21, 22–24 asexual multiplication (merogony), 16, 18, 19, 20 cell invasion and internalization, 13–16, 17, 18 environmental stage and infection source, 12 excystation, 13 extracellular stages, 20–21 prepatent patent times, 19–20 sexual reproduction (gamogony), 16–17, 21, 22, 23 sporogony, 18–19, 24 prevention, control, and treatment, 28–35 chemicals reducing oocyst viability, 31–33, 34–35 environmental factors reducing oocyst numbers, 33, 35
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physical factors reducing oocyst viability, 28–31 structure, 21–26, 27 attachment for feeder organelle, 14, 15, 23–24 oocyst, 21–22, 24, 25, 26 sexual stages, 21, 22, 23, 25–26 sporozoites, 17, 18, 22–23 trophozoites and merozoites, 17, 19, 24–25, 27 taxonomy, 4–10 genotypes, 8 mammalian host species, 5–6 transmission, 26, 28 Biophysical methods, oocyst concentration from feces, 174–183 centrifugation, 178–179 enteropathogenic parasite screen, Cryptosporidium requests as part of, 182 flotation of oocysts, 182–183 formol-ether concentration, oocysts, 182 formol-ether concentration for oocysts, 179 infection in absence of detectable oocysts, 183 purification of oocysts, 183 Biopsy, 193–194, 391 Birds, 395–413 avian cryptosporidiosis, 396–397 avian genotypes, 410–413 duck genotype, 412 Eurasian woodcock, 412 goose genotypes, 412–413 types I-IV, 410–412 chemoprophylaxis and control, 399 chemotherapy and prophylaxis, 276 Cryptosporidium baileyi, 10, 403–407 biology, 403, 404 genetic analysis, 407 host range, 406–407 life cycle, 403, 405–406 pathogenesis, 406 Cryptosporidium galli, 407–410 biology, 407–409 genetic analysis, 410 host range, 410 life cycle, 409 pathogenesis, 409–410 Cryptosporidium meleagridis, 10, 399–403 biology, 399–400 discovery of, 2 genetic analysis, 403 host range, 401–402 life cycle, 400 pathogenesis, 400–401 domesticated fowl diagnostic methods, 175–176 differences among species infecting humans and agricultural animals, 178 evolutionary relationships of avian isolates, 404 host specificity, 10 immunity, 398–399 molecular epidemiology human infections, 134 role of animals in transmission to humans, 139
species and genotypes, 130–132 transmission routes in animals, 132 respiratory disease in, 256 species infecting, 11 in vitro and in ovo culture, 398–399 Bison, 471–472 Bleach, 399, 503, 505, 510, 511, 512, 514, 518 Blood samples, seroepidemiology/population immunity, 85 Bovine colostrum, 223–224, 451 chemotherapy and prophylaxis, 273 human volunteer studies, 98 protective effect, 223 in reptiles (snakes), 391 Bovine genotype, 9 hosts infected by, 11 nomenclature, 121 Bovine serum albumin (BSA), 195, 223 Breast feeding, 86, 223, 238 Bright-field microscopy, 176 Brine, oocyst flotation, 182, 183 Buffalo, 5, 420, 421, 471
C Calcium carbonate flocculation, oocyst concentration from water, 311–312 Calves animal models of disease, 487 chemotherapy and prophylaxis, 274–275 dairy cattle manure treatment within calf facilities, 379–380 differences among species infecting humans and agricultural animals, 178 experimental generation of C. parvum oocysts, 486–487 hyperimmune bovine colostrum and, 224 molecular epidemiology, 132 role of animals in transmission to humans, 138, 139 subtype families, 84 neonatal diarrhea, 80 oocyst shedding, species, 379 parasite propagation and oocyst production, calves, 507 pathogenic species, 175 prepatent and patent period, 19 serial passage and loss of virulence, 454 source of infection in pigs, 467 species infecting, 461, 462 transmission to rabbits, 424 Camel, 5, 392, 421, 426, 429, 471 chemotherapy and prophylaxis, 275 species infecting, 11 Canids, 5 Captive species, 429 Carbohydrate metabolism, 46, 59–61 Carbol fuchsin, 186 Carcass decontamination, food processing, 94 Carnivora, 5
52268_Index.fm Page 531 Thursday, September 27, 2007 10:38 AM
Index Carnivores, 421–422 Carrier state, see also Asymptomatic/subclinical infection snakes, 389 Case surveillance studies, recreational water outbreaks, 339–341 Cation pumps, 46, 68 Cats, see also Cryptosporidium felis chemotherapy and prophylaxis, 276 C. muris experimental transmission to, 421 feral, 421 latent infection, prednisolone treatment, 151 prepatent and patent period, 19, 20 species infecting, 11 Cattle, 3, 428, 452–462, 463, 464 animal models for generation of C. parvum oocysts, 486–487, 507 chemotherapy and prophylaxis, 274–275 clinical disease and pathology, 452–454 detection and identification, 454 early studies, 2 interleukin-12 treatment, 219 molecular epidemiology C. hominis in, 83, 84 epidemic human cryptosporidiosis, 145 MboII for RFLP, 166 as source of infection in surface water, 147 transmission routes, 132–133 neonatal diarrhea, 80 pathogenic species, 175 prepatent and patent period, 19, 20 prevalence, 455–458, 459–460, 461, 462 age of animal and, 461, 462 beef cattle, nonmolecular methods, 457 dairy cattle, nonmolecular methods, 456 mixed dairy and beef cattle, nonmolecular methods, 458 primate-human-cattle transmission cycle, 425 species and genotypes, 452, 453, 454–455 age of animal and, 461, 462 beef cattle, molecular methods, 460 dairy cattle, molecular methods, 459–460 mixed dairy and beef cattle, molecular methods, 461 subtyping, 458, 460–461, 463–464 waste management beef cattle manure, 381 dairy cattle manure, 378–381 CD4 and CD8 positive T-cells, 214–215, 216, 217, 225 CD40, 214 Cell culture, see In vitro systems/cell culture Cell culture PCR, genotyping Cryptosporidium in natural water, 148 Cell-free culture systems, Cryptosporidium behavior in, 255 Cell invasion and internalization, life cycle, 13–16, 17, 18 Cell lines, 500–505, 515 Cell-mediated immune response, see also T-cell mediated immunity birds, 397–398
531 elimination of infection, 86 Cellulose filtration, oocyst purification, 509 Centers for Disease Control, 344 outbreaks in U.S., 341 water park use survey, 338 Centrifugation, oocyst, 178–179, 506 gradient, see Gradient purification water sample concentration, 311, 312 Cervine genotype(s), 8, 429 as emerging human pathogen, 430 epidemiology, 83 evolutionary relationships of avian isolates, 404 food contamination, 292 hosts infected by, 11 host specificity, 11 in humans, 175 laboratory lemurs, 427 molecular epidemiology, 132 epidemic human cryptosporidiosis, 144 genotyping Cryptosporidium in natural water, 148 human infections, 134, 136, 138 neighbor-joining analysis of SSU rRNA gene, 131 RFLP, 167 role of animals in transmission to humans, 138 ruminants, 421 sheep, 465 source tracking, 146–147 wastewater effluent oocysts, 373 wild mammals, 430 Cesium chloride purification, oocyst, 177, 183, 509, 510, 513 CGS-A, CGS-B, CGS-C, 429 Chaperons, 58 Chemical disinfectants, 31–33, 34–35; see also Chlorine dioxide/chlorination; Ozone treatment Chemokines, 211; see also Cytokines Chemotherapy and prophylaxis, 255–279 animal models for testing, 485, 488–491 gerbil, 490–491 gnotobiotic pig, 489–490 immunosuppressed rodent, 489 neonatal mouse, 488–489 SIV-infected rhesus monkey, 490 in animals, 265–279 miscellaneous, 276 rodents, 266–273 ruminants, 274–275 in vitro, 277–278 birds, 399 in humans, 256–265 clinical manifestations, 256 future directions, 265 HAART, 263–264 supportive therapy, 264–265 in humans, agents for, 256–263 benzeneacetonitrile derivatives, 261–262 macrolides, 257–261 miscellaneous, 262–263 rifabutin (Mycobutin®), 261
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livestock, 451–452 Chicken egg yolk antibody, 268, 277 Chickens C. baileyi, 27 C. baileyi antigens, effects on embryos, 224 chemotherapy and prophylaxis, 273, 276 chorioallantoic membrane culture, 500 immunized, antibodies from, 224 prepatent and patent period, 19, 20 species infecting, 11 Children, see also Infants and children chemotherapy and prophylaxis, 258 clinical disease and pathology, 237–238 molecular epidemiology C. parvum versus C.hominis, 146 endemic human cryptosporidiosis, 140, 141 epidemic human cryptosporidiosis, 141–142 human infections, 135 transmission routes dogs as source of infection, 441 recreational water contamination, 337, 339 wild mammals, 430 Chipmunk genotype, 138, 146–147 Chiroptera, 5 Chlorine dioxide/chlorination, 32, 33 recreational water disinfection, 358 wastewater effluent treatment, 374 water disinfection, 321–322 Cholangiopathy, AIDS, 240 Chorioallantoic membrane (CAM) culture, 500 C. baileyi, 398–399 in vitro methods, 501, 519, 520 Chromatography, 190, 511 Chromosome 6, 45, 128 Chromosomes, 44 Chronic infections, 1 Cladogram, 49 Clarifiers, waste management, 375 Clarithromycin, 258, 259, 261, 267, 272, 277 Clinacox® (diclazuril), 261–262 Clinical disease and pathology, 234–244 animal models, 487–488 cattle, 452–454 C. baileyi, 403 C. galli, 409–410 chemotherapy and prophylaxis in humans, 256 histopathology, 240–241 human infectivity trials, 242–244 life cycle, see Life cycle life cycle and, see Life cycle manifestations, 235–239 age-related infections, 236, 237–239 age-related infections, children, 237–238 age-related infections, elderly, 238–239 in immunocompetent host, 235–236 in immunocompromised host, 236–238 molecular epidemiology, 150 C. parvum versus C.hominis, 146 differences among Cryptosporidium species in human cryptosporidiosis, 145–146 utility of tools, 127
organ sites, 239–240 extraintestinal sites, 239–240 GI tract, 239–240 pathophysiology, 241–242 pigs, 466, 467–468 reptiles, 389–390 symptom duration C. parvum versus C.hominis, 146 epidemiology, 81–82 Clinical infections, infectivity trials, 243 Cloacal swabs, diagnosis in reptiles, 390 Clonal population structure, 128, 129, 130 Coccidia, 2, 4, 22, 23, 25, 28, 57, 58, 69, 261, 399 chemotherapy, 255, 261, 263, 265 life cycle, 6 oocyst structure, 19 phylogenetic relationships, 49, 50, 51 in vitro systems, 500, 514, 518, 520 Coding capacity, genome properties, 46 Coinfection, see Mixed infections/coinfection Colchicine, 277 Colostral antibodies, 225, 226 Colostrum, see Bovine colostrum Commercial kits and reagents, diagnostic, analytic, nucleic acid purification antibodies, 184, 185 diagnostic, 180–181 DNA extraction, 164, 165, 195 enzyme immunoassays, 189 immunomagnetic separation (IMS), 164 oocyst preparations, 183 Common shrew, 424 Communities population genetics, significance of, 130 utility of tools, 127 Companion animals, 437–444 cats, 437–439 dogs, 440–441, 442 horses, 442–444 transmission routes, 26, 139 Comparative genomics, 49–51, 52 Complement, 211 Composting treatment beef cattle manure, 381 dairy cattle manure, 380 sludge, 377, 378 Computer programs oocyst detection, laser scanning cytometry, 313–314 population genetics, 169 Concurrent infections, see Mixed infections/coinfection Conserved sequences multilocus sequence analysis, 127 and specificity of PCR, 122 Contamination population genetics, significance of, 130 source tracking, 146–151 utility of tools, 127 Continuous flow (CF) centrifuge, 312
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Index Control and prevention, see Prevention, control, and treatment Copy number, target gene interspecies differences versus intragenotypic variation, 7 and RFLP, 122 sequence differences between copies, 123 and SSCP method, 125 Council of State and Territorial Epidemiologists (CSTE), 341 COWP, see Cryptosporidium oocyst wall protein (COWP) Coyote, 11 Coyote genotype, 8 CPB-DIAGF/R, 197–198, 199, 200 Cpgp40/15, see GP60 Cross-reactivity antibody response, 85–86 Johnson et al. primers, 197–198 Crosstransmission, see also Transmission early studies, 9 molecular epidemiology, 132 wildlife to domestic animals and humans, 419 Cryopreservation, oocyst, 506 CryptoDB, 59 Cryptosporidium andersoni in cattle, 452, 453 age and, 457 clinical disease and pathology, 454 molecular analysis, 455 species and genotypes, 461 species and genotypes by age, 462 species and genotypes in dairy cattle, 459, 460 species and genotypes infecting beef cattle, 460 dairy waste, 379 diagnosis in humans, 175 in livestock and domesticated fowl, 175 morphology, 175 PCR, comparison of methods, 199 PCR-RFLP, 200 evolutionary relationships of avian isolates, 404, 410 genetic similarities at SSU rRNA locus, 412 hosts infected by, 11 host specificity, 11 life cycle, 20 molecular epidemiology age of animal and, 132 DdeI for RFLP, 166 genotyping Cryptosporidium in natural water, 148, 149 genotyping tools, 122 neighbor-joining analysis of SSU rRNA gene, 131 RFLP, 167 role of animals in transmission to humans, 138 source tracking, 147 in sheep, 465 sites of infection, 11 terminology caveats, 121
533 valid species, host types, and oocyst measurements, 9 wastewater effluent oocysts, 373 in wildlife, 427, 428, 429 Cryptosporidium anserinum, 396 Cryptosporidium baileyi antibody protective effect, 223 in birds, 396, 403–407 biology, 403, 404 genetic analysis, 407 host range, 406–407 life cycle, 403, 405–406 pathogenesis, 406 chemotherapy and prophylaxis, 273, 276 chorioallantoic membrane culture, 520 developmental stage comparison, 401 diagnosis commercial oocyst preparations, 183 differences among species infecting humans and agricultural animals, 178 in livestock and domesticated fowl, 175 PCR, comparison of methods, 199 evolutionary relationships of avian isolates, 404 genetic similarities at SSU rRNA locus, 407 genomics, phylogenetic analysis, 50 hosts infected by, 11 host specificity, 10 immune response, 397–398 life cycle, 27 molecular epidemiology genotyping Cryptosporidium in natural water, 148, 149 neighbor-joining analysis of SSU rRNA gene, 131 RFLP, 167 role of animals in transmission to humans, 138 source tracking, 147 oocyst detection in shellfish, 296 oocyst morphometry, 400 prepatent and patent period, 19, 20 in reptiles (snakes), 392 sites of infection, 11 valid species, host types, and oocyst measurements, 9 virus coinfection, 397 in vitro and in ovo culture, 398–399 wastewater effluent oocysts, 373 Cryptosporidium blagburni, 409 Cryptosporidium bovis in cattle, 453 age and, 457, 462 molecular analysis, 455 species and genotypes, 461 species and genotypes by age, 462 species and genotypes in dairy cattle, 459, 460 dairy waste, 379 diagnosis differences among species infecting humans and agricultural animals, 178 molecular methods, 200 hosts infected by, 11
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molecular epidemiology age of animal and, 132 genotyping, 132 neighbor-joining analysis of SSU rRNA gene, 131 RFLP, 167 role of animals in transmission to humans, 138 prepatent and patent period, 20 in sheep, 465 valid and nonvalid species, 7–8 valid species, host types, and oocyst measurements, 9 in wildlife, 427 Cryptosporidium bovis-like genotype, sheep, 465 Cryptosporidium canis in cattle, 453, 455 diagnosis differences among species infecting humans and agricultural animals, 178 in humans, 175 MAS-PCR, failure to detect in human samples, 201 PCR, comparison of methods, 199 PCR-RFLP, 197, 200 in dogs, 441, 442 epidemiology, 93 evolutionary relationships of avian isolates, 404 genotypes, 8 host range, 430 hosts infected by, 11 molecular epidemiology C. parvum versus C.hominis, 146 endemic human cryptosporidiosis, 141 genotyping, 132 genotyping Cryptosporidium in natural water, 148, 149 genotyping tools, 122 human infections, 134, 135, 136–137 neighbor-joining analysis of SSU rRNA gene, 131 RFLP, 167 role of animals in transmission to humans, 138, 139 source tracking, 147 subtyping tool limitations, 125 waterborne outbreaks, 126 valid and nonvalid species, 7–8 valid species, host types, and oocyst measurements, 9 in wildlife, 427, 428, 429, 430 Cryptosporidium felis in cats, 437–439 in cattle, 453, 455 diagnosis differences among species infecting humans and agricultural animals, 178 ELISA-based serodiagnostic assay, 193 in humans, 175 MAS-PCR, failure to detect in human samples, 201 PCR, comparison of methods, 199
PCR, nested, 198–199 PCR-RFLP, 197, 200 epidemiology, 83 IgG antibody response to reference strain of C. parvum, 85–86 percentage of human cases caused by, 83 zoonotic transmission, 93 evolutionary relationships of avian isolates, 404 host range, 11, 430 host specificity, 11 molecular epidemiology endemic human cryptosporidiosis, 141 genotyping Cryptosporidium in natural water, 149 genotyping tools, 122 human infections, 134, 135, 136–137 latent infection, 151 neighbor-joining analysis of SSU rRNA gene, 131 RFLP, 167 role of animals in transmission to humans, 138, 139 source tracking, 147 subtyping tool limitations, 125 waterborne outbreaks, 126 prepatent and patent period, 19, 20 valid species, host types, and oocyst measurements, 9 in wildlife, 427 Cryptosporidium galli birds, 396, 407–410 biology, 407–409 genetic analysis, 410 host range, 410 life cycle, 409 pathogenesis, 409–410 differences among species infecting humans and agricultural animals, 178 evolutionary relationships of avian isolates, 404 genetic similarities at SSU rRNA locus, 407, 412 genotypes, 8 hosts infected by, 11 in livestock and domesticated fowl, 175 morphology, 175 neighbor-joining analysis of SSU rRNA gene, 131 oocyte morphometry, 400 RFLP, 167 transmission to humans, 138 valid and nonvalid species, 7–8 valid species, host types, and oocyst measurements, 9 Cryptosporidium hominis, see also specific topics biochemical studies, lack of, 58 in cattle, 84, 453 molecular analysis, 455 species and genotypes, 461 C. meleagridis phylogenetic relationships, 403 diagnosis differences among species infecting humans and agricultural animals, 178 ELISA-based serodiagnostic assay, 193
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Index fluorescence staining intensity, 189 in humans, 175 PCR, 198–199 PCR-RFLP, 197, 200 epidemiology, 84 geographic distribution, 95 percentage of human cases caused by, 82, 83 risk factors, 87 transmission routes, 90 evolutionary relationships of avian isolates, 404 genome sequences and metabolic pathway maps, 59 genomics, 45, 46, 47 hosts infected by, 11 molecular epidemiology, 150 behavior in human versus C. parvum, 146 in cattle, 84 endemic human cryptosporidiosis, 139–140, 141 epidemic human cryptosporidiosis, 141–145 genotypes and subtypes, 121 genotyping, 132 genotyping Cryptosporidium in natural water, 148, 149 GP60, 124 host specificity, 145 human infections, 134, 135, 136–137 neighbor-joining analysis of SSU rRNA gene, 131 population genetics, 129–130 RFLP, 167 role of animals in transmission to humans, 139 source tracking, 147 subtype families, 84 subtyping, PCR-sequencing analysis of GP60, 167–169 subtyping tools, 125 waterborne outbreaks, 126 oocyst generation in piglets, 487 in pigs, 466, 467 in sheep, 465 TU502, 45 valid and nonvalid species, 7–8 valid species, host types, and oocyst measurements, 9 wastewater effluent oocysts, 373 waterborne outbreaks, 344 in wildlife, 427 Cryptosporidium meleagridis in birds, 396, 399–403 biology, 399–400 genetic analysis, 403 host range, 401–402 life cycle, 400 pathogenesis, 400–401 in vitro and in ovo culture, 398–399 developmental stage comparison, 401 diagnosis differences among species infecting humans and agricultural animals, 178 in domesticated fowl, 175
535 ELISA-based serodiagnostic assay, 193 in humans, 175 PCR, comparison of methods, 199 PCR, nested, 198–199 PCR-RFLP, 197, 200 discovery of, 2 in dogs, 441, 442 epidemiology IgG antibody response to reference strain of C. parvum, 85–86 percentage of human cases caused by, 82, 83 zoonotic transmission, 93 evolutionary relationships of avian isolates, 404 genetic similarities at SSU rRNA locus, 407 host range, 11, 430 host specificity, 10 molecular epidemiology, 150 endemic human cryptosporidiosis, 141 genotyping, 164 genotyping Cryptosporidium in natural water, 148, 149 genotyping tools, 122 human infections, 134, 135, 136–137 neighbor-joining analysis of SSU rRNA gene, 131 RFLP, 167 role of animals in transmission to humans, 138 subtyping tools, 125 waterborne outbreaks, 126 oocyte morphometry, 400 in pigs, 467 recreational water use and, 341 in reptiles (snakes), 392 valid species, host types, and oocyst measurements, 9 waterborne outbreaks, 350 in wildlife, 427 Cryptosporidium molnari, 387–388 hosts infected by, 11 valid species, host types, and oocyst measurements, 9 Cryptosporidium muris in cats, 438, 439 in cattle, 452 diagnostic methods, 174 commercial oocyst preparations, 183 differences among species infecting humans and agricultural animals, 178 in humans, 175 MAS-PCR, failure to detect in human samples, 201 morphology, 175 PCR, 199 PCR-RFLP, 197, 200 epidemiology, percentage of human cases caused by, 83 evolutionary relationships of avian isolates, 404, 410 genetic similarities at SSU rRNA locus, 412 genomics, 47, 50 genotypes, 8
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host range, 430 hosts infected by, 11 immune response B-cell mediated, 223 cytokines and infection control, 219 T-cell mediated immunity, 213, 215 in laboratory animals, 427 life cycle, 11 molecular epidemiology DdeI for RFLP, 166 genotyping in natural water, 148, 149 genotyping tools, 122 human infections, 134 RFLP, 167 role of animals in transmission to humans, 138 source tracking, 147 subtyping tool limitations, 125 terminology caveats, 121 phylogenetic analysis, 50 in pigs, 470, 471 prepatent and patent period, 19 in reptiles (snakes), 392 in rodents, 423 in ruminants, 421 sites of infection, 11 valid and nonvalid species, 7–8 valid species, host types, and oocyst measurements, 9 wastewater effluent oocysts, 373 in wildlife, 427, 428, 429, 430 in zoo animals, 426 Cryptosporidium oocyst wall protein (COWP), 82 biochemistry, 66–67 cattle species and genotypes, 461 C. baileyi, 407 C. meleagridis, 403 coding capacity, 46 diagnostic methods, 200 mixed infections, 201 PCR, primers and step cycle protocols, 197, 198 functional genomic analysis, 47 molecular epidemiology genotyping Cryptosporidium in natural water, 149 genotyping tools, 122 human infections, 134 source tracking, 147 typing and subtyping methods, 202 Cryptosporidium parvum, see also specific topics in amphibians, 389 in cats, 438 in cattle, 452, 453 age and, 457 clinical features and pathology, 453–454 molecular analysis, 455 species and genotypes, 461 species and genotypes by age, 462 species and genotypes in dairy cattle, 459, 460 species and genotypes infecting beef cattle, 460 subtyping, 458, 460–461, 463–464 chorioallantoic membrane culture, 520
dairy waste, 379 diagnosis commercially available antibodies, 185 differences among species infecting humans and agricultural animals, 178 ELISA-based serodiagnostic assay, 193 fluorescence staining intensity, 189 in humans, 175 in livestock and domesticated fowl, 175 PCR, 198–199 PCR-RFLP, 197, 200 discovery of, 2 in dogs, 441, 442 epidemiology geographic distribution, 95 human volunteer studies, 98 IgG antibody response to reference strain of, 85–86 percentage of human cases caused by, 82, 83 risk factors, 87 transmission routes, 89 zoonotic transmission, 93 evolutionary relationships of avian isolates, 404 genome sequences and metabolic pathway maps, 59 genomics comparison of genome annotation, 45 "omics" resources, 47 orthologous gene clusters across cladogram, 49 phylogenetic analysis, 50 physical mapping, 44 in gorillas, 425 in horses, 444 host range, 11, 430 host specificity, 10–11 IOWA type 2, 45 molecular epidemiology, 150 anthroponotic versus zoonotic subtypes, 124 behavior in human versus C. hominis, 146 endemic human cryptosporidiosis, 139–140 epidemic human cryptosporidiosis, 141–145 genotypes and subtypes, 121 genotyping Cryptosporidium in natural water, 148, 149 GP60, 124 host specificity, 145 human infections, 134, 135, 136–137 neighbor-joining analysis of SSU rRNA gene, 131 population genetics, 128–129 population genetics, significance of, 130 RFLP, 167 role of animals in transmission to humans, 138, 139 source tracking, 147 subtype families, 84 subtyping, PCR-sequencing analysis of GP60, 167–169 subtyping tools, 125 waterborne outbreaks, 126 oocyst generation in calves, 486–487
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Index phylogenetic relationships, C. meleagridis, 403 in pigs, 466, 467, 468, 471 prepatent and patent period, 19 in ruminants, 421 in sheep, 465 sites of infection, 11 transmission, 26–27 transmission from calves to rabbits, 424 valid and nonvalid species, 7–8 valid species, host types, and oocyst measurements, 9 wastewater effluent oocysts, 373 waterborne outbreaks, 344 in wildlife, 427, 428, 429 Cryptosporidium pestis, 83, 84, 121 Cryptosporidium rhesi, 425 Cryptosporidium saurophilum, see Cryptosporidium varanii (saurophilum) Cryptosporidium scophthalmi, 387–388 hosts infected by, 11 valid species, host types, and oocyst measurements, 9 Cryptosporidium serpentis, 131, 167, 199, 200 in amphibians, 389 avian Cryptosporidium species evolutionary relationships, 404 genetic similarities, 410, 412 chemotherapy and prophylaxis, 273 genomics, phylogenetic analysis, 50 hosts infected by, 11 in reptiles (snakes), 389, 392 sites of infection, 11 valid species, host types, and oocyst measurements, 9 Cryptosporidium suis in cattle, 453 species and genotypes by age, 462 species and genotypes infecting beef cattle, 460 diagnosis differences among species infecting humans and agricultural animals, 178 in humans, 175 MAS-PCR, failure to detect in human samples, 201 PCR, comparison of methods, 199 PCR-RFLP, 197, 200 epidemiology, percentage of human cases caused by, 83 evolutionary relationships of avian isolates, 404 host range, 430 hosts infected by, 11 host specificity, 11 manure treatment, 381 molecular epidemiology age of animal and, 133 genotyping, 132 neighbor-joining analysis of SSU rRNA gene, 131 RFLP, 167 role of animals in transmission to humans, 138 subtyping tool limitations, 125
537 pigs, 467, 468, 470, 471 prepatent and patent period, 19, 20 sheep, 465 valid and nonvalid species, 7–8 valid species, host types, and oocyst measurements, 9 in wildlife, 427 in wild mammals, 430 Cryptosporidium suis-like genotype, cattle, 459 Cryptosporidium tyzzeri, 396 Cryptosporidium varanii (saurophilum), 199, 200 evolutionary relationships of avian isolates, 404 hosts infected by, 11 neighbor-joining analysis of SSU rRNA gene, 131 nomenclature, 121 in reptiles (snakes), 389 RFLP, 167 valid species, host types, and oocyst measurements, 9 Cryptosporidium wrairi differences among species infecting humans and agricultural animals, 178 evolutionary relationships of avian isolates, 404 hosts infected by, 11 host specificity, 11 molecular epidemiology, RFLP, 167 neighbor-joining analysis of SSU rRNA gene, 131 PCR, comparison of methods, 199 PCR-RFLP, CPB-DIAGF/R primers, 200 phylogenetic relationships, C. meleagridis, 403 in reptiles (snakes), 392 valid species, host types, and oocyst measurements, 9 Crystal structures, proteins, 47 CSL glycoprotein, see Life cycle Culture conditions, 515–516 Cyclospora, 6, 182, 186, 291 Cytokines, 217–222, 225 chemotherapy and prophylaxis, in animals, 265 other proinflammatory cytokines, 220–221 parasite killing, 221–222 pathophysiology, 241, 242 Th1, 217–219 Th2, 219–220 Cytometry, 312–314 Cytopathic effect, 242 Cytoskeleton, 66, 70, 241, 242 CZB genotype, 83
D Dairy cattle C. andersoni, 452 manure, waste management, 378–381 prevalence, 455, 457 prevalence, nonmolecular methods, 456, 458 species and genotypes, molecular methods, 459–460, 461 DAPI staining, 189, 313, 314, 513, 517 Databases
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genomes and metabolism, 59 genomics, 52–53 “-omics” resources, 47, 48 Day-care outbreaks, 237 DDBJ, 48 DdeI, 164, 166, 200 DE-52 chromatography, sporozoite purification, 511 Deer, 5, 11, 426 Deer genotype, 8, 11 Deer-like genotype, 8 in cattle, 453 age and, 457, 462 dairy waste, 379, 459 molecular analysis, 455 species and genotypes, 459, 462 hosts infected by, 11 molecular epidemiology age of animal and, 132 RFLP, 167 Deermouse genotype, 8, 404 Defensin-beta, 211 Delayed hypersensitivity (DH), birds, 398 Dendritic cells, 212, 219 Detection limits, oocyst concentration from feces, 178, 179 Developmental stages, C. meleagridis and C. baileyi, 401 DFMO (difluoromethylornithine), 263 DHFR, see Dihydrofolate reductase (DHFR) Diagnostic index, nested PCR, 183 Diagnostic methods, 174–203 antibody detection, 191–193 enzyme-linked immunoelectrotransfer blot (EITB), 191–192 enzyme-linked immunosorbent assay, 192–193 biopsy, 193–194 cattle, 454 immunological methods, antigen detection, 188–190 enzyme-linked antibodies, 189–190 fluorescence-labeled antibodies, 188–189 laboratory diagnosis rationale, 176–178 molecular, nucleic acid detection methods, 194–203 DNA extraction, 194–197 primer, gene locus selection, and PCR, 197–203 oocyst concentration from feces, 178–184 biophysical methods, 174–183 immunomagnetic separation (IMS), 183–184 oocyst morphology, 175–176 oocyst staining methods, 186–188 auramine phenol, 186–187 Ziehl-Neelsen stain, modified, 187–188 outbreak and large scale epidemiological investigations, 184–185 pathogenic species, 174–175 in humans, 175 in livestock and domesticated fowl, 175–176 pigs, 468 reporting results of PCR RFLP/sequencing, 203 reptiles, 390–391 sensitivity of detection in feces, 190–191
shipping of oocysts and oocyst DNA for QA and round robin testing, 203 Diagnostics, population genetics and, 130 Diaper-aged children, recreational water contamination, 337 Diaper changing, 87, 88, 93, 95, 97 Diclazuril (Clinacox®), 261–262 Differential interference contrast (DIC) microscopy, 176, 313, 314–315, 513, 518 Difluoromethylornithine [(DFMO) Eflornithine®, Ornidyl®], 263 Dihydrofolate reductase (DHFR), 70 nucleotide metabolism, 62 PCR, 147, 197 source tracking, 61 Dihydrofolate reductase-thymidylate synthase (DHFR-TS), 62 Diphenoxylate, 264 Direct oocyst smear, 177 Disease burden, epidemiology, 96–97 Disease tracking, PCR, 202–203 Disinfection, environment, see Prevention, control, and treatment Disinfection, water, 320–325 artificial recreational water bodies, 357 chlorine dioxide, 321–322 deficiencies, 352–357 ozone, 321 prevention of outbreaks, 358–360 regulatory requirements, 306 ultraviolet light, 322–325 Divergent proteins, drug targets, 70 Diversity, genetic, see Genetic diversity Diversity, host, 10; see also Host range DNA immunization with, 224, 226 polyclonal antibodies from immunization with, 224 ribosomal small subunit gene, see SSU 18S rRNA gene toll-like receptor activation, 211 DNA extraction from fecal samples using QIAamp kit, 164 molecular diagnostic methods, 194–197 outbreak and large scale epidemiological investigations, 194–195 small numbers of partially purified oocysts, 195–197 oocyst detection in shellfish, 296 from water and fecal samples using FastDNA SPIN Soil kit, 165–167 from water samples using QIAamp kit, 165 DNA hybridization, 295 DNA metabolism, 58–59 DNA repair, 30, 68–69 DNA sequencing, see Sequencing DnaSP, 169 Dog(s), 5; see also Cryptosporidium canis C. meleagridis infection, 401 C. muris experimental transmission to, 421 role of animals in transmission to humans, 139
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Index species infecting, 11 Dog genotype, 8, 11 Domain architectures, surface proteins, 49, 50 Domestic animals C. muris experimental transmission to, 421 companion animals, 437–444 Dot blots, 454 Double-stranded RNA, 69 birds, genetic analysis, 403 C. meleagridis, 403 diagnostic methods, 294 subtyping tools, 125, 197, 202 DraI, 199 Drinking water-related outbreaks/exposure, 305–325 epidemic human cryptosporidiosis, 142–143, 145 epidemiology, 88 burden of disease, 96–97 incubation period, 80 intervention, 99 protection of vulnerable populations, 95 seasonality, 95–96 transmission routes, C. hominis, 90 transmission routes, C. parvum, 89 oocyst recovery methods, 309–316 complete methods, 314–315 detection methods, 313–314 monitoring, data quality, 315–316 sample collection, 309–311 sample concentration, 311–312 separation from sample debris, 312–313 prevalence of Cryptosporidium in water, 306–309 early studies, 306–308 recent studies, 308–309 regulatory requirements, 316–318 United Kingdom, 316–317 United States, 317–318 seroepidemiology/population immunity, 85 treatment, 318–325 disinfection, 320–325 filtration, 318–320 Drought, and waterborne outbreaks, 91 Drug targets, 62 biochemical divergence from other apicomplexans and, 57 potential, 69–70 Drying, and oocyst viability, 30 Duck genotype, 8, 412 evolutionary relationships of avian isolates, 404 GenBank accession numbers, 411 genetic similarities at SSU rRNA locus, 407 hosts infected by, 11 molecular epidemiology genotyping, 132 RFLP, 167 Ducks, 11 Duration of infection, 1; see also Time course of infections Duration of symptoms C. parvum versus C.hominis, 146 epidemiology, 81–82 Dynabeads, 164
539
E Effluent treatment, waste management, 372–375 Eflornithine® (difluoromethylornithine), 263 Eggs, see Chorioallantoic membrane (CAM) culture Egg yolk antibody, 268, 277 18S rRNA, see SSU 18S rRNA gene Eimeria, 2, 45, 131, 261, 295, 404 biochemistry apicoplast, 63 fatty acid metabolism, 63, 64 mannitol cycle, 61 polyamine metabolism, 65 expressed sequence tags, 44 oocyst structure, 19 taxonomy, 4, 6 tritiated uracil incorporation, 518 in vitro cultivation, 514 Elderly, clinical disease and pathology, 238–239 Electrolytes cation pumps, 46, 68 management of, 265 pathophysiology, 241–242 Electron microscopy, 193 Electrophoresis, PCR product, 196 EMBL, 48 Endemic disease C. hominis population genetics, 129 genotype and subtype identification for, 121 human cryptosporidiosis, 139–141 molecular epidemiology, 150 transmission routes, 120 Endoscopy, diagnosis in reptiles, 390 Energy metabolism, 59–61 amylopectin and, 28 coding capacity, 46 Enrofloxacin, 270, 399 Enterocyte cell lines, 211, 220, 221, 222, 277 Enterocytes, 11, 80, 224 asexual reproduction in, 183 avian Cryptosporidium infections Cryptosporidium baileyi, 403, 405 duck genotype, 412 biopsy, 193 cat, 421 histopathology, 240 host defense, 224 complement production, 212 cytokines and, 222, 225 IL-1 and reproduction in, 221 intraepithelial lymphocytes, 216 nitric oxide synthase activity, 221 NK cytotoxicity, 213 immune response, cytokine production, 241 innate immunity, 211–212 sodium pump, 241 Enteropathogenic parasite screen, Cryptosporidium requests as part of, 182 Environmental chemicals, oocyst sensitivity, 98 Environmental contamination, wildlife species, 420
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Environmental controls, see Prevention, control, and treatment Environmental exposure, epidemiology, 89 Environmental factors and human infections, 134 and oocyst viability, 33, 35 and waterborne outbreaks factors contributing to transmission in recreational waters, 336 rainfall, see Rainfall Environmental samples analysis of, 125–126 commercial diagnostic kits, 180–181 DNA extraction from water samples, 164–165 epidemic human cryptosporidiosis, 142–143 genotyping by PCR/RFLP analysis of SSU rRNA gene, 165–167 subtyping by PCR sequencing analysis of GP60 gene, 167–169 Environmental stage and infection source, life cycle, 12 Enzyme immunoassays cultured parasite development, 518 horses, 443 Enzyme-linked immunoassays antigen detection, 189–190 commercial diagnostic kits, 180–181 comparison of diagnostic methods, 191 diagnostic methods, 189–190 Enzyme-linked immunoelectrotransfer blot (EITB), 191–192 Enzyme-linked immunosorbent assays (ELISA), 85, 190 antibody detection, 192–193 cattle, 454 commercial diagnostic kits, 180–181 dogs, 440–441 genotyping tools, 122 pigs, 468 reptiles (snakes), 391 Epidemic disease C. hominis population genetics, 129 C. parvum population genetics, 128–129 genotype and subtype identification for, 121 human cryptosporidiosis, 141–145 Epidemic population structure, 130 Epidemiology, 80–100 age and gender, 86, 87 age and susceptibility, 86 asymptomatic infection, 81 descriptive and analytical investigations, 97 diagnostic methods, 184–185 MAS-PCR, 200–201 PCR, 202–203 differences in Cryptosporidium species, 87 disease burden, 96–97 duration of symptoms, 81–82 enzyme-linked immunoelectrotransfer blot (EITB), 192 exclusions on basis of infection, 95 geographic distribution, 95
historical developments, 3 human volunteer studies, 98 immunocompromised populations, 86 incubation period, 80–81 infants and children, 86 intervention, 99 mixed infections, 98 molecular, 85–86; see also Molecular epidemiology molecular methods, 3–4 oocyst viability, infectivity, quality, and ID50, 98 period prevalence, 98 population genetics and, 130 protection of vulnerable populations, 95 recreational water outbreaks, 339–341 risk assessment, 99 risk factors, 87, 88–89 seasonality, 95–96 seroepidemiology/population immunity, 85–86 sources, 87 sporadic disease, 97 surveillance, 80 transmission routes, 92, 93 drinking water, 90–92 food and drink, 93–94 other drinking waters, 92 person-to-person, 93 recreational waters, 92–93 travel, 94 zoonotic, 93 Epifluorescence, 314 Epithelial barrier function, 242 Epithelial cells, 211–212; see also Enterocytes Epitope-paratope interaction, immunomagnetic separation (IMS), 184 EST, see Expressed sequence tags (ESTs) Ether, 179 Ethyl acetate, 179 Eurasian woodcock genotype, 407, 411, 412 Evans Blue, 189 Evolutionary/phylogenetic relationships, 6, 255 animal cryptosporidiosis, 131 avian isolates, 404 biochemistry carbohydrate and energy metabolism, 61 divergence from other apicomplexans, 58 C. meleagridis, 403 C. molnari, 388 genetic analysis, 407 genomics, 49–51, 52 interspecies differences versus intragenotypic variation, 7, 120 molecular epidemiology, 120, 124, 130, 131, 138 mouse genotype, 429 nucleotide pathways, 52 oocyst structure and, 21 with sarcocystids or calyptosporids, 21 Excretion of oocysts, see Oocyst shedding Excystation, 13, 28 Expressed sequence tags (ESTs), 44, 45, 47 Expression vectors, cryptosporidial proteins, 70–71 Extracellular stages, life cycle, 20–21
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Index Extrachromosomal double-stranded RNA, 202 Extraintestinal sites, clinical disease and pathology, 239–240
F Farm animals, see Cattle; Livestock/agricultural animals; Pig(s); Sheep Farms, source tracking, 147 FastDNA SPIN Soil kit, 164, 165–167 Fatty acid metabolism, 63–65 Fatty acid transporters, 46 Fecal excretion of oocysts, see Oocyst shedding Fecal samples, see Stool specimens Feeder organelle attachment, 14, 15, 23–24 Feral pigs, 468 Ferret genotype, 8, 199, 200 evolutionary relationships of avian isolates, 404 neighbor-joining analysis of SSU rRNA gene, 131 Ficoll gradients, 509 Filters water sample concentration, 311–312 water sampling, 309–311 Filtration, oocyst purification, 509 Filtration of water, 96–97, 99, 318–320 artificial recreational water bodies, 357 bag or cartridge, 320 conventional, 319 diatomaceous earth, 320 direct, 319 membrane, 320 recreational water disinfection, 358 regulatory requirements, 306 slow sand, 319–320 wastewater effluent, 372, 375 Fish, 387–388 host specificity, 10, 11 species and genotypes, 11, 130–132 Fixation/fixatives, 190, 194 Flies, transmission routes, 88, 94 Flocculation, oocyst concentration from water, 311–312 Flotation, oocysts, 178, 182–183, 194, 500, 509, 510 environmental sample analysis, 126 fecal samples, 178 oocyst recovery from water sample debris, 312 recovery, 182–183 water sample debris, 312 Flotation, sporozoites, 511 Flow cytometry, 312–313, 314, 519 Flow filtration, oocyst concentration from water, 312 Fluids and electrolytes management of, 265 pathophysiology, 241–242 Fluorescein isothiocyanate (FITC) staining, 177, 185, 186, 187, 188, 189, 190, 296, 313, 314 Fluorescein thiosemicarbazide (FTSC) labeling, 512, 513, 514, 517, 518, 519 Fluorescence labeling, 177, 182 antigen detection, 188–189
541 commercial diagnostic kits, 180–181 commercially available antibodies, 185 FITC-labeled mABs, 177, 186, 188 in vitro studies, 512, 513 in vitro studies, flow cytometry, 519 Foals, 444 Foodborne disease, 289–300 commercial diagnostic kits, 180–181 confirmed outbreaks, 291–293 detection methods, 293–297 beverages, oocysts in, 295 fresh fruits, vegetables, or prepared foods, 293–295 shellfish, 295–297 epidemiology sources, 88 transmission routes, 89, 90, 93–94 Hazard Analysis and Critical Control Points (HAACP) and other management regulations, 299 inactivation, 298–299 molecular epidemiology, 120, 150 epidemic human cryptosporidiosis, 141, 142–143, 145 sources of contamination, 138 subtyping tools, 144, 145 utility of tools, 127 possible contamination sources, 297–298 transmission routes, 3 Foodborne Diseases Active Surveillance Network (FoodNet), 97 Food handlers, 88, 292, 299 epidemic human cryptosporidiosis, 145 foodborne disease sources, 297 Food processing, foodborne disease sources, 297 Formalin, 176, 179, 194 Formol-ether concentration, oocysts, 178, 179, 182, 183, 184–185, 194 41kD antigen, seroepidemiology, 85 Fountains, 355–356 Fowl, see also Birds; Chickens diagnostic methods, 175–176 differences among species infecting humans and agricultural animals, 178 Fox(es), 5, 11, 422, 429 Fox genotype(s), 8, 11, 404, 429, 430 Fragment typing and subtyping methods, 202 Freeze-thaw cycles, and oocyst viability, 28 Freezing, and oocyst viability, 28, 29 Fresh vegetables, see Foodborne disease Fresh water, see also Waterborne outbreaks deficiencies, 351–352 prevention of outbreaks, 357–358 Functional analysis, genome properties, 47 Fusion proteins, 70–71
G γ/δ receptor positive T-cells, 214 Gamma irradiation, oocysts, 29, 225
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Gamogony (sexual reproduction), 16–17, 21, 22, 23 Gamont-like stages, C. andersoni, 20 Gamonts, 513 Gastric lavage, diagnosis in reptiles, 390 Gastrointestinal tract clinical disease and pathology, 239–240 histopathology, 240–241 life cycle, see Life cycle sites of infection, see Site of infection Gel electrophoresis PCR product, 196 surface proteins, species identification, 7 Gene clusters, 49 Gene copies interspecies differences versus intragenotypic variation, 7, 125 and RFLP, 122 sequence differences between copies, 123 and SSCP method, 125 Gene length, 45 General biology, see Biology Gene silencing, RNA-mediated, lack of, 69 Genetic analysis, see also specific techniques avian Cryptosporidium species, 411, 412 avian genotypes I-IV, 410 C. baileyi, 407 C. galli, 410 C. meleagridis, 403 evolutionary relationships of, 404 genetic similarities, 407 Cryptosporidium taxonomy, 120 molecular epidemiology, 82, 84; see also Molecular epidemiology species identification, 7 wildlife and zoo mammals, 429–430 Genetic diversity animal infections, 133 wild mammals, 427–428 Genetics, animal models, 491–492 Gene transfer, interspecies/horizontal, 51, 52, 127 Genome annotation, 45, 46–47, 59, 60, 61 Genome project, Cryptosporidium parvum, 59 Genome survey sequences, 44–45 Genomics, 4, 43–53 biochemical divergence from other apicomplexans, 57–58 comparative, 48–52 horizontal (interspecies) gene transfer, 51, 52 phylogenetics, 49–51 databases, 47, 48 gene regulation, 48 genome properties, 45–47 annotation, 45, 46–47 coding capacity, 46 functional analysis, 47 genome resources, 48 molecular data types, 44–45 prospects, 51–53 Genotype 1, see Cryptosporidium hominis Genotype 2, see Cryptosporidium parvum Genotypes, 8
avian Cryptosporidium species, 411 cattle, 452, 453, 454–455 age of animal and, 461, 462 beef cattle, molecular methods, 460 dairy cattle, molecular methods, 459–460 mixed dairy and beef cattle, molecular methods, 461 diagnostics number of, 175 typing and subtyping methods, 202 interspecies differences versus intragenotypic variation, 7 livestock cattle, see Genotypes, cattle molecular analysis, 455, 457 pigs, 470, 471 molecular epidemiology, 82, 84, 120–121, 150 animal cryptosporidiosis, 131 endemic human cryptosporidiosis, 141 host range, 84 human cryptosporidiosis, 133–138 in mammals, birds, reptiles, and fish, 130–132 waterborne outbreaks, 126 nomenclature, 84–85 number of, 2 taxonomy, 8 valid and nonvalid species, 7–8 wastewater effluent oocysts, 373 wild mammals, 428–429 Genotyping, 122, 123 endemic human cryptosporidiosis, 141 natural water samples, 148–149 oocysts in fecal and environmental water samples, 164–169 source tracking, 147 Genus, 1 Genus-specific subtyping tools, 122–123 Geographic distribution cattle, 453 epidemiology, 95 descriptive and analytical investigations, 97 seasonality, 95–96 human infections, 12 livestock, see Prevalence; specific types of livestock (cattle, sheep, etc.) molecular epidemiology, 84, 150 animal transmission routes, 133 C. parvum population genetics, 129 endemic human cryptosporidiosis, 139–140, 141 human infections, 135, 136–137 population genetics, 127 utility of tools, 126 prevalence of Cryptosporidium in water, 306–309 early studies, 306–308 recent studies, 308–309 Gerbils animal models, 146, 490–491 chemotherapy and prophylaxis, 269 Giemsa stain, 174, 186 Glutamine, 259
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Index Glycine, 259 Gnotobiotic pig, 145–146, 466, 489–490 Goats, 465–466 chemotherapy and prophylaxis, 275 epidemiology C. hominis occurrence, 83 C. parvum in small mammals and, 83 host species, 5 immunization, 224 species infecting, 11 Goose, 11 Goose genotype(s), 8, 396, 412–413 evolutionary relationships of avian isolates, 404 GenBank accession numbers, 411 genetic similarities at SSU rRNA locus, 407 hosts infected by, 11 molecular epidemiology neighbor-joining analysis of SSU rRNA gene, 131 role of animals in transmission to humans, 138, 139 RFLP, 167 Gorillas, 425 GP60, 164 biochemistry, 67 C. meleagridis, 403 C. parvum subtypes in cattle, 463–464 molecular epidemiology, 82 animal transmission routes, 133 C. hominis population genetics, 130 C. parvum population genetics, 129 endemic human cryptosporidiosis, 140 epidemic human cryptosporidiosis, 142–143, 144, 145 MLST primers, 170–171 PCR sequencing analysis, 167–169 subtype families, 84 subtyping, 83 subtyping tools, 123, 124, 125 typing and subtyping methods, 202 Gradient purification, 183, 509, 511, 512, 513 environmental sample analysis, 126 oocyst recovery from water sample debris, 312 Green monkeys, 425 GRO-alpha, 211 Guinea pigs, 3, 11, 421
H HAART therapy, 86, 256, 259, 260, 265, 275 chemotherapy and prophylaxis, 263–264 molecular epidemiology, 150 Halofuginone, 265 Halogen disinfectants, 32, 33; see also Chlorine dioxide/chlorination HAPPY map technique, 44, 45 Hazard Analysis and Critical Control Points (HAACP), 299 Heat shock protein-70, see HSP-70 Heat shock proteins, 47
543 Hedgehog, 427 Hepatobiliary system involvement, 241 Heterogeneity, genetic C. hominis population genetics, 129, 130 endemic human cryptosporidiosis, 140 multilocus sequence analysis, 127–128 Highly active antiretroviral therapy (HAART), see HAART therapy High-pressure methods, oocyst inactivation, 299 Histology, 193 Histopathology clinical disease and pathology, 240–241 pigs, 467 HIV/AIDS, 258 biliary tract involvement, 239–240 chemotherapy and prophylaxis, 255, 256–257, 260–261 clinical disease in, 236–237 clinical manifestations with, 256 epidemiology, 80, 86 incubation period, 81 protection of vulnerable populations, 95 transmission routes, 93 molecular epidemiology, 150, 151 endemic human cryptosporidiosis, 141 human infections, 134 role of animals in transmission to humans, 139 pathophysiology, 242 SIV-infected rhesus monkey model, 490 transmission routes, wild mammals, 430 Horizontal (interspecies) gene transfer, 46, 47, 51, 52, 127 drug targets, 70 protein domain architecture, 49 Horse(s), 3, 5, 11, 132 Horse genotype, 8, 444 hosts infected by, 11 molecular epidemiology, 132 Hospital infection, 88, 89, 90 Host, adaptation to and cross-transmission, 430 molecular epidemiology, 83 transmission routes in animals, 132 Host-adapted genotypes, wild mammals, 427–428 Host diversity, 10 Host factors and course of infection, 1–2 molecular epidemiology, 150 population genetics, 127 Host range animal models and, 493 avian Cryptosporidium species, 396–397 C. baileyi, 406–407 C. galli, 410 C. meleagridis, 401–402 Cryptosporidium species, 178 molecular epidemiology, 83, 84 adaptation by types to host species, 83 human infections, 134 role of animals in transmission to humans, 138 transmission routes in animals, 132–133
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species identification, 7 valid Cryptosporidium species, 9 Host species, 1, 5–6 Host specificity, 83 animal models and, 493 biology, 7–10 C. parvum versus C. hominis, 145 Cryptosporidium taxonomy, 120 geographic distribution of human cryptosporidiosis, 12 species identification, 7 Host susceptibility, 127 Hot springs, 351–352 Hot water, oocyst sensitivity, 94 HSP 65, 58 HSP-70, 58 avian Cryptosporidium species, 411 C. baileyi, 407 C. galli, 410 cattle, 460, 461 diagnostic methods, 197, 202 epidemiology, 82 genotyping tools, 122 molecular epidemiology C. hominis population genetics, 129 C. parvum population genetics, 129 genotyping Cryptosporidium in natural water, 148 MLST primers, 170–171 subtyping tools, 125 PCR, 197 PCR primers, 121 pigs, 471 typing and subtyping methods, 202 Human cryptosporidiosis chemotherapy and prophylaxis, see Chemotherapy and prophylaxis C. meleagridis infection, 402 diagnostic methods, 175 differences among species infecting humans and agricultural animals, 178 epidemiology species causing, 82–83 strain differences, 87 geographic distribution, 11 host specificity, 10 molecular epidemiology, 133–146 biological, clinical, and epidemiologic differences among Cryptosporidium species, 145–146 C. parvum and C. hominis as most common species infecting, 84 C. parvum population genetics, 128–129 endemic, 139–141 epidemic, 141–145 as source of infection in surface water, 147 species and genotypes, 133–138 subtype families, 84 prepatent and patent period, 19–20 species causing, 11, 82–83, 178 transmission routes, 26
animal Cryptosporidia infecting, 10 from horses, 444 primate cross-transmission, 425 wildlife as source, 419–420 Human-specific isolates, C. parvum, 83 Human (anthroponotic) transmission, 3, 26, 120 molecular epidemiology, 150 C. parvum phylogenetic relationships, 124 endemic human cryptosporidiosis, 140 mother-to-infant, 151 population genetics, significance of, 130 primate cross-transmission, 425 recreational waters, see Recreational water to reptiles (experimental), 392 Human trials/volunteer studies, 236 epidemiology, 98 infectivity, 242–244 Humatin® (paromomycin), 258, 259, 260, 262 Husbandry practices, 139–140 Hybridization, water samples, 314 Hydrogen peroxide, 32, 34 Hygiene and sanitation, see Prevention, control, and treatment Hyperimmune bovine colostrum, 223–224, 451 chemotherapy and prophylaxis, 273 human volunteer studies, 98 in reptiles (snakes), 391 Hypochlorite, 399, 503, 505, 510, 511, 512, 514, 518
I ID50, 98 Identification cattle, 454 molecular epidemiology, utility of tools, 126 Immune responses, 210–226 animal models, 487–488 antibodies, parasite-specific, 222–224 immunization-induced, 223–225 infection-induced, 222–223 and course of infection, 1–2 cytokines and infection control, 217–222 other proinflammatory cytokines, 220–221 parasite killing, 221–222 Th1, 217–219 Th2, 219–220 innate immunity, 210–213 components, 210 epithelial cells, 211–212 natural killer cells, 212 T-cell mediated immunity, 213–217 importance of T cells in infection control, 213–215 mucosal compartments, 215–216 parasite antigen-specific priming of T-cells and T-cell memory, 216–217 vaccination against infection, 225 Immune status, host, 1–2 Immunity birds, 398–399
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Index epidemiology asymptomatic infection, 81 endemic human cryptosporidiosis, 141 human volunteer studies, 98 incubation period, 81 and oocyst negative stool samples, 183 population - seroepidemiology, 85–86 oocyst downregulation, 183 and severity of infection, 215, 223, 256 Immunization-induced antibodies, parasite-specific, 223–225 Immunoblotting, 85 Immunochromatography, 180–181, 190, 191 Immunocompromised host(s), see also HIV/AIDS asexual reproduction of cryptosporidia in enterocytes, 183 asymptomatic infection in, 243 chemotherapy and prophylaxis, 258 clinical disease and pathology, 236–238 and course of infection, 2 dogs and, 441 epidemiology, 82, 84, 86, 88 molecular epidemiology, 84, 134, 150 pathophysiology, 241 Immunodiagnostic methods antibody detection, 191–193 antigen detection, 188–190 commercial diagnostic kits, 180–181 cultured parasite development, 518 detection of oocysts in food, 294–295 dogs, 440–441 horses, 443 immunomagnetic separation (IMS), 183–184 sensitivity of detection, 190–191 water samples, 314 Immunofluorescence assays, environmental sample analysis, 126 Immunofluorescence staining, 513 cattle, 454 commercial diagnostic kits, 180–181 comparison of diagnostic methods, 191 developing parasites in culture, 516–517 horse samples, 443 oocyst detection in shellfish, 296 Immunoglobulins, see also Antibodies; Immunity human volunteer studies, 98 subtypes secretory IgA, 222–223 seroepidemiology, 85 time course of infection, 191–192 time course of infection, 222 Immunomagnetic separation (IMS), 194 commercial diagnostic kits, 180–181 commercial kit, 164 diagnostic methods, oocyst concentration from feces, 183–184 genotyping Cryptosporidium in natural water, 148, 149 molecular epidemiology, 126, 148, 149 oocyst detection in shellfish, 296 pig samples, 468
545 Immunomodulators, human volunteer studies, 98 Immunosuppressed Mongolian gerbil, 146 Immunosuppressed rodent models, 489 Imodium, 264 Inactivation, foodborne Cryptosporidia, 298–299 Incidence period prevalence, 98 seasonality, 95–96 Incident control team (ICT), 97 Incubation period, 242 epidemiology, 80–81 in humans, 175 Indinavir, 259, 260 Indirect immunofluorescence methods, 85 Indomethacin, 399 Infants and children, see also Children epidemiology, 86 diaper changing, transmission via, 87, 88, 93, 95, 97 exclusions on basis of infection, 95 interpersonal transmission, 93 molecular, 150, 151 mother-to-infant transmission, 151 previous infection and age of children as factor in antibody levels, 86 recreational water as route of transmission, 340 sources, 88 zoonotic transmission, 93 recreational water contamination, 337–338 Infection in absence of detectable oocysts, 183 Infection-induced antibodies, parasite-specific, 222–223 Infective dose cattle, 453 epidemiology, 89 human volunteer studies, 98 incubation period, 81 infectivity trials, 242–243 molecular epidemiology, 150 oocyst, 98 Infectivity, 7 human trials, 242–244 in vitro, age of oocyst and, 511 in vitro assays, 314 molecular epidemiology pathogenic species, 147, 149 role of animals in transmission to humans, 138 oocyst, 98, 511 prevention, control, and treatment measures and, 28 species identification, 7 sporozoite, 511 water samples, 314 Inflammatory mediators, 211; see also Cytokines Information Collection Rule (ICR), 308, 314, 315 Inhibitors, polymerase chain reaction (PCR) environmental sample analysis, 125–126 stool specimens, 194–195 Taq polymerase, reagents affecting, 196 Innate immunity, 211–212, 225 Inoculum density, in vitro systems, 514–515
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Inoculum size, 81; see also Infective dose Inosine monophosphate dehydrogenase (IMPDH), 51, 62 In ovo culture, see also Chorioallantoic membrane (CAM) culture avian Cryptosporidia, 398–399 chorioallantoic membrane culture, ?? method, 520 Insectivora, 5 Insectivores, 424 Insects food contamination, 298 transmission routes, 88, 94 In situ hybridization, 314 Interferon-gamma, 217–218, 219, 220, 225, 241 Interleukin-1, 221 Interleukin-2, 218–219 Interleukin-4, 219–220, 225 Interleukin-5, 220 Interleukin-6, 221 Interleukin-8, 211, 242 Interleukin-12, 212, 214, 218–219 chemotherapy and prophylaxis, 258 Interleukin-13, 220 Interleukin-15, 213, 221 Interleukin-18, 221 Internalization, cell, 13–16, 17, 18 International Code of Zoological Nomenclature, 83, 120, 121 Interpersonal transmission, 120 elderly, 239 epidemiology, 88, 93 C. hominis, 90 C. parvum, 89 sporadic disease, 97 molecular epidemiology, 151 Interspecies differences, intraspecies variability and, 7 Interspecies (horizontal) gene transfer, 46, 47, 51, 52, 127 drug targets, 70 protein domain architecture, 49 Interspecific recombination, 69, 127 Intervention, 84, 99 Intracellular development stage cattle, 453 lack of RNA from, 47 Intracellular gene transfer, 51 Intracellular localization, oocysts, 193 Intraepithelial lymphocytes, 216 Intragenotypic variation, see Subgenotypes; Subtypes Intraspecies variability, versus interspecies differences, 7 Introns, 44, 45, 46, 69 Invasion and internalization, life cycle, 13–16, 17, 18 In vitro systems/cell culture, 500–520 animal models versus, 492 avian embryos, 520 birds, 398–399 chemotherapy studies, 277–278 chronology of developments, 500–505 culture, 511–520
host cells (cell lines) and environment, 515, 516 oocyst and sporozoite preparation, 511–514 quantification of developing parasites, 516–519 disinfection testing, 31 drug efficacy tests, 277–278 epithelial integrity, 242 infectivity assays, 314 interferon-gamma, 222 parasite production and purification, 505–511 oocyst purification, 507–510 oocyst sources and storage conditions, 505–507 sporozoite excystation, 510 sporozoite purification, 510–511 stool collection, 507 Ionic conditions, and immunomagnetic separation, 184 Ion pumps, 46, 68 Iron-sulfur proteins, 58 Irradiation, 299 Isolate identification/discrimination C. parvum, human-specific, 83 interspecies differences versus intragenotypic variation, 7 intragenotypic variation versus different copies of gene, 122 methods for, 120, 121 molecular epidemiology, 82 Isolates, with complete genome sequences, 127 Isospora, 4, 19, 21, 182, 186, 437, 467 ITS-1/ITS-2, 122–123, 125
J Japanese field mouse genotype, 8 Jenner’s stain, 186 Johnson et al. primers, 197–198 Juveniles, 132
K Kangaroo genotype, 199, 200 Kangaroos, 5, 424, 429 Kaopectate, 264 Karyotype, 44 Kidneys, C. baileyi, 406 Koala genotype, 199, 200 Kyoto Encyclopedia of Genes and Genomes (KEGG), 48, 59
L Laboratory animals C. muris experimental transmission to, 421 experimental models, see Animal models rodents, 428 Lactobacilli, 259, 269 Lagomorphs, 5, 423–424 Lagoon treatments dairy waste, 379–380
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Index sludge, 377 swine manure, 382 Lakes, 351–352 Lambs, 138, 139 Lamina propria, 216, 241 Lamivudine, 259, 260 Lasalocid, 265 Laser scanning cytometry, 313–314 Latent infection, 151 Legislation, drinking water treatment, 99 Lemurs, 424, 427 Letrazuril, 258, 262 Life cycle, 9, 11–21, 22–24 apicomplexans, 4, 6 asexual multiplication (merogony), 16, 18, 19, 20 avian Cryptosporidium species C. baileyi, 403, 405–406 C. galli, 409 C. meleagridis, 400 cell invasion and internalization, 13–16, 17, 18 environmental stage and infection source, 12 excystation, 13 extracellular stages, 20–21 morphology, 7 prepatent patent times, 19–20 sexual reproduction (gamogony), 16–17, 21, 22, 23 species identification, 7 sporogony, 18–19, 24 taxonomic classification of cryptosporidia, 4 in vitro studies, 513 Light microscopy, see Microscopic detection Linkage disequilibrium, 128, 129 Livestock/agricultural animals cattle, 452–462, 463, 464; see also Cattle clinical disease and pathology, 452–454 detection and identification, 454 prevalence, 455–458, 459–460, 461, 462 species and genotypes, 452, 453, 454–455 subtyping, 458, 460–461, 463–464 diagnostic methods, 175–176 differences among species infecting humans and agricultural animals, 178 goats, 465–466 host species, 5 miscellaneous, 471–472 molecular epidemiology, 84 animal transmission routes, 132, 133 C. parvum population genetics, 128–129 endemic human cryptosporidiosis, 139–140 pigs, 466–471 ELISA-based assays and PCR, 468 microscopic detection, 469–470 molecular methods for species and genotype identification, 471 sheep, 462, 464, 465; see also Sheep sporadic disease, 97 transmission routes, 3, 26 primate-human-cattle transmission cycle, 425 small mammals as source of reinfection, 83 watershed contamination, 429–430 wild animals, cross transmission, 420
547 Live vaccines, 225 Lizard genotype, 8, 11, 131 Lizards, 11, 121 Llamas, 471 Lomotil, 264 Loperamide (Imodium), 264 Lymph nodes, parasite antigen in, 215 Lymphocytes, mucosal compartments, 215–216
M Macaques, 6, 424–425 Macrogamonts, 16, 17, 26 Macrolides, 257–261 Macrophages, 212 Mammals host species, 5–6 molecular epidemiology avian pathogen infectivity, 134 species and genotypes, 130–132 Mannose-binding lectin, 212 Manure, see also Livestock/agricultural animals food contamination, 298 waste management, see Waste management Marine water, 350–351, 357–358 Marmosets, 5, 424 Marsupial genotype(s), 8, 404, 429, 465 Marsupials, 424 host species, 5 molecular epidemiology genotyping, 132 neighbor-joining analysis of SSU rRNA gene, 131 RFLP, 167 MAS-PCR, 200–201 Maternal antibodies, 86 Maximum likelihood analysis, 410 MboII, 164, 166 MCP-1, 211 Media, culture, 516 Meiosis, 17 Meiotic recombination, 69 Melting curve analysis, 122, 197 Membrane filtration of water, 96–97, 99, 309–311, 320 Membrane proteins and transporters, 67–68 Mepron® (atovaquone), 263 Merogony (asexual multiplication), 16, 18, 19, 20, 24 Meront, 513 Meronts, 16 Merozoites, 16, 513 attachment for feeder organelle, 23 structure, 17, 19, 24–25, 27 Metabolism, see also Biochemistry biochemical divergence from other apicomplexans, 57–58 drug targets, 57, 69–70 highly streamlined, 58–59 MICA, 213 MICB, 213 Microarrays, 122, 202
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Microbial risk assessments (MRAs), 99 Microgametes, 22 attachment for feeder organelle, 23–24 sexual stages, 25–26 Microgamonts, 16, 17, 21, 25 Micrometry, 176 Microsatellite analysis, see Minisatellite/microsatellite analysis Microscopic detection, 193, 313 cattle, 455 C. galli, 408 cultured parasite development, 516–518 diagnostic methods, in cattle, 454 oocyst morphology, 176 pigs, 468, 469–470 sheep, 464 Microtubules, 66 Microwave radiation, and oocyst viability, 29 Minisatellite/microsatellite analysis, 82 complete genome sequences and, 127–128 molecular epidemiology MLST of C. parvum, C. hominis, and C. meleagridis, 169 MLST primers, 170–171 subtyping tools, 123, 125 typing and subtyping methods, 202 Minor species mixed infections, detection methods, 123 nomenclature, 85 MIP-2alpha, 211 Mirazid, 258 Mitochondria, absence of, 45, 46, 58 Mitochondria, relict, 18 Mitochondrial origin of nuclear genes, 46 Mixed infections/coinfection birds, 397, 398 cattle, species and genotypes in dairy cattle, 459 C. meleagridis and C. baileyi, 401 diagnostic methods, PCR, 201 epidemiology, 98 molecular epidemiology, 84, 150 animal infection with mixed C. parvum subtypes, 133 genotyping, 164 genotyping C. in natural water, 149 human infections, 136–137 PCR limitations, 123 population genetics, 127, 130 pigs, 467, 468 Molecular diagnostic methods, 194–203 DNA extraction, 194–197 primer, gene locus selection, and PCR, 197–203 Molecular epidemiology, 85–86, 120–151, 164–171 animal cryptosporidiosis, 130–133 species and genotypes in mammals, birds, reptiles, and fish, 130–132 transmission routes, 132–133 contamination source tracking, 146–151 genotyping and subtyping oocysts in fecal and environmental water samples, 164–169 DNA extraction, 164-165
genotyping by PCR/RFLP analysis of SSU rRNA gene, 165–167 subtyping by PCR sequencing analysis of GP60 gene, 167–169 human cryptosporidiosis, 133–146 biological, clinical, and epidemiologic differences among Cryptosporidium species, 145–146 endemic, 139–141 epidemic, 141–145 species and genotypes, 133–138 multilocus sequence typing of C. parvum, C. hominis, and C. meleagridis, 169–171 population genetics, 127–130 C. hominis, 129–130 C. parvum, 128–129 MLT and MLST, 127–128 significance of in diagnostics and epidemiology, 130 tools and methods, 120–127 concepts for species, genotypes, and subtypes, 120–121 environmental sample analysis, 125–126 genotyping, 122, 123 subtyping, 123–125 utility of, 126–127 wild mammals, 427–429 genetic diversity, 427–428 species and genotypes, 428–429 Molecular methods, 3–4, 7 cultured parasite development, 518 historical developments, 3–4 species and genotype identification in cattle, 455, 457 in pigs, 471 Mollusks, see Shellfish Mongolian gerbil, immunosuppressed, 146 Mongoose genotype, 8 Monkey(s), 3 host species, 5, 6 laboratory animals, 427 SIV-infected rhesus monkey model, 488, 490 Monkey genotype, 8, 199 in humans, 175 neighbor-joining analysis of SSU rRNA gene, 131 PCR-RFLP, CPB-DIAGF/R primers, 200 RFLP, 167 Monoclonal antibodies chemotherapy and prophylaxis in animals, 265 commercially available, 185 FITC-labeled, 186, 188 immunomagnetic separation (IMS), 184 oocyst and sporozoite immunolabeling, 517 passive immunization with, 224 in vitro studies flow cytometry, 519 life cycle, 513 Morphology, 6–7, 175–176 Morphometry, oocyst, 120, 175, 176, 178 avian Cryptosporidium species and genotypes, 400 cattle, 452
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Index Mortality, 86 Mosaicism, interspecific, 52, 127 Mother-to-infant transmission, 151 Mouse, 422, 423, 428 animal models, 488–489 chemotherapy and prophylaxis, 266–269 C. muris experimental transmission to, 421 host species, 6 oocyst production in, 506–507 prepatent and patent period, 19 serial passage and virulence, 454 species infecting, 11 wildlife genotypes, 429 Mouse genotype, 9, 199, 428–429 evolutionary relationships of avian isolates, 404 hosts infected by, 11 molecular epidemiology genotyping Cryptosporidium in natural water, 148 neighbor-joining analysis of SSU rRNA gene, 131 role of animals in transmission to humans, 138 wastewater effluent oocysts, 373 Mucosal compartments, T-cell mediated immunity, 215–216 Mucosal immunization, 226 Multilocus typing (MLT) and multilocus sequence typing (MLST), 82 C. meleagridis, 403 molecular epidemiology, 127–128, 164 C. hominis population genetics, 129 of C. parvum, C. hominis, and C. meleagridis, 169–171 C. parvum population genetics, 128–129 population genetics, 127 population genetics, 127–128 primers, 170–171 Multiple episodes of infection, endemic human cryptosporidiosis, 141 Multiple species, infections with, see Mixed infections/coinfection Multiplex allele-specific PCR (MAS-PCR), 200–201 Multiplex bead assay, 85 Multiplexing real-time polymerase chain reaction (PCR), 197 Multi-species infections, see Mixed infections/coinfection Muskrat genotype(s), 8, 11, 404, 429 Muskrats, 11, 167, 423, 429 Mycobutin® (rifabutin), 261
N Natural killer cells, 212 Natural swimming areas, circulation in, 357 Nature reserves, 420 NCBI GenBank, 48 Neighbor-joining analysis, 131, 404 Neonatal calves, 80, 456 experimental models, 487
549 oocyst production in, 505–506, 507 Neonatal goats, 466 Neonatal mouse model, 324, 488–489, 492 immune response hyperimmune bovine colostrum and, 223 IL-4 role, 219 IL-12 effects on oocyst challenge, 218 interferon expression, 217–218 passive protection with mABs, 224 passive protection with sIgA, 223 T-cell mediated immunity, 214 vaccination, 225 oocyst production in, 506–507 Neonatal sheep, 462, 488 Neonates, 151, 213 chemotherapy and prophylaxis, 279 drug effectiveness, 266, 267 ELISA-based serodiagnostic assay, 193 oocyst sources, 505 Nested polymerase chain reaction (PCR) 18S rRNA gene fragments, 197, 198–199 Cryptosporidium oocyst wall protein (COWP), 200 genotyping tools, 122 molecular epidemiology genotyping, 164 genotyping Cryptosporidium in natural water, 148, 149 sensitivity of detection, 147 source tracking, 147 oocyst negative stool samples, 183 typing and subtyping methods, 202 water samples, 314 New genotypes and species, see Novel (new) species/genotypes NF-κB, 211, 242 Nichols-Johnson 18S rRNA assay, 198, 199 Nitazoxanide (Alinia®), 70, 259, 263 drug targets, 70 livestock, 451 Nitric oxide and derivatives, 221 Nomenclature, 83, 84 Cryptosporidium taxonomy, 120 historical developments, 2–3 molecular epidemiology, 121 three prime repeat region polymorphisms and, 125 valid and nonvalid species, 7–8 Nonhuman primates, 424–426 Non-nucleoside reverse transcriptase inhibitors (NNRTIs), 263, 264 Nonspecific amplification, primers and, 122 Nosocomial infection, 88 Novel (undetermined) species/genotype, 83 Novel (new) species/genotypes, 7, 9, 10, 452, 471 birds, 412 companion animals dog genotype, C. canis, 441 horse genotype, 444 diagnosis, 174 genotyping Cryptosporidium in natural water, 148, 149 Japanese field mouse genotype, 428
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livestock, 461, 465 methods for identification of, 120, 121 nomenclature, 50 percentage of human cases caused by, 83 swine manure, 381 Novel subtype emergence, 130 Nuclear genome, genes of mitochondrial origin in, 46 Nucleic acid detection, 194–203 DNA extraction, 194–197 primer, gene locus selection, and PCR, 197–203 Nucleic acid/nucleotide metabolism, 58–59, 62 horizontal (interspecies) gene transfer, 51 salvage pathways, absence of biosynthetic pathways, 46, 51 Nucleic Acid Sequence Based Amplification (NASBA), 314 Nucleoside reverse transcriptase inhibitors (NRTIs), 263, 264 Nucleotides, coding, 45 Nutritional status children, 237–238 C. parvum versus C.hominis, 146 and infection rates, 86 infection sequelae, 86, 146 mice, and cytokine-mediated parasite killing, 221 molecular epidemiology, 146, 150
O Occupational exposure, epidemiology, 88 Octreotide (Sandostatin), 264 Omnivores, 421 Oocyst negative stool samples, 183 Oocysts animal systems for production of, 486–487 cattle C. andersoni, 455 dairy cattle manure applied to soil, 380–381 purification of, 454 chorioallantoic membrane culture, 520 commercially available antibodies, 185 dairy cattle manure applied to soil, 380–381 detection methods, 294–295 diagnostic methods, see also Diagnostic methods concentration from feces, 178–184 morphology, 175–176 staining methods, 186–188Ooo DNA and RNA metabolism, 68–69 drinking water, recovery from, 309–316 complete methods, 314–315 detection methods, 313–314 monitoring, data quality, 315–316 sample collection, 309–311 sample concentration, 311–312 separation from sample debris, 312–313 epidemiology asymptomatic shedders, 81 detection in foods, 94 human volunteer studies, 98
molecular methods for identification of source, 82 sources, 87 viability, infectivity, quality, and ID50, 98 gamma-irradiated, vaccination with, 225 life cycle, see Life cycle morphology, see Morphology recovery, concentration, and purification of from cattle, 454 diagnostic methods, 178–184 from drinking water, 311–313 flowchart, 508 from food, 293–294 for in vitro studies, 507–510 removal from drinking water, 96–97 seroepidemiology, 85 size measurement, see Morphometry, oocyst species identification, 6–7 structure, 21–22, 24, 25, 26 taxonomic classification of cryptosporidia, 4 transmission routes, 3 in vitro systems, see In vitro systems/cell culture wall protein, see Cryptosporidium oocyst wall protein (COWP) waste management, see Waste management Oocyst shedding asymptomatic shedders, 81 C. meleagridis and C. baileyi, 401 C. parvum versus C.hominis, 146 dogs, 441 human volunteer studies, 98 immunization and, 224 infectivity trials clinical versus subclinical infections, 243 time course of infection, 242 livestock cattle, 452 pigs, 468 sheep, 462 recreational water contamination, 337 time course of infection, 256 wild mammals versus domestic animals, 429–430; see also specific wild animals Oocyst size C. andersoni, 455 C. curyi, 8 valid Cryptosporidium species, 9 Opossum, 167, 424 Opossum genotype(s), 8, 404, 429 Organ sites, clinical disease and pathology, 239–240 extraintestinal sites, 239–240 GI tract, 239–240 Ornidyl® (difluoromethylornithine), 263 Ortholog clusters, 49 Ostrich genotype, 8 Outbreak control team (OCT), 97 Outbreaks day-care, 237 diagnostic methods, 184–185 molecular epidemiology epidemic human cryptosporidiosis, 141–145
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Index population genetics, significance of, 130 recreational water-related, see Recreational water Ovine genotype, 8 Oxidative metabolism, 46 Ozone treatment, 99 oocyst sensitivity, 299 wastewater effluent, 372, 374 water disinfection, 321
P Panmictic population structure, 129 Parasite screen, Cryptosporidium requests as part of, 182 Paromomycin (Humatin®), 70 animals, 265, 273 birds, 399 calf, 274 cat, 276 chicken, 270, 273 fish, 391 gerbil, 269 goat, 275 lamb, 275 livestock, 399, 451 mice, 266, 267, 268, 489 piglet, 490 rats, 270, 271 ruminants, 265 drug targets, 70 humans, 258, 259, 260, 262 in vitro studies, 277, 278 Parturition, 151, 462 Passive immunization, 223, 225–226; see also Bovine colostrum with monoclonal antibodies, 224 in reptiles (snakes), 391 Patent period infectivity trials, 242 pigs, 467 Pathogenesis C. baileyi, 406 C. galli, 409–410 C. meleagridis, 400–401 Pathogenicity, 1 C. parvum versus C.hominis, 146 loss of, serial passage and, 453–454 Pathogenic species, diagnostic methods, 174–175 in humans, 175 in livestock and domesticated fowl, 175–176 Pathology, 127 birds, 397 histopathology, 240–241 organ sites, 239–240 extraintestinal sites, 239–240 GI tract, 239–240 pathophysiology, 241–242 reptiles, 389–390 Pathophysiology animal models, 487–488
551 birds, 397 clinical disease and pathology, 241–242 Peptides, antimicrobial, 211 Percoll gradients, 183, 509, 511, 513 Period prevalence, 98 Perissodactyla, 5 Person-to-person transmission, see Interpersonal transmission Pest infestations, foodborne disease sources, 297 Pet food, contamination of, 93 Peyer’s patches, 215 pH and immunomagnetic separation, 184 oocyst sensitivity, 98 sludge treatment, 377 Phase contrast microscopy, 176 Phylogenetic analysis, 6, 7 cat isolates, 438 Cryptosporidia in wild animals, 427, 428 goose genotypes, 412 Phylogenetic relationships, see Evolutionary/phylogenetic relationships Physical factors reducing oocyst viability, 28–31; see also Ultraviolet light Physical genome mapping, 44 Pig(s), 3, 466–471 animal models for research, 487–488, 489–490 chemotherapy and prophylaxis, 276 ELISA-based assays and PCR, 468 experimental generation of C. hominis oocysts, 487 feral, 421, 468 gnotobiotic, 145–146, 466, 489–490 host species, 5 microscopic detection, 469–470 molecular methods for species and genotype identification, 471 prepatent and patent period, 20 species infecting, 11 waste management, manure treatment, 381–382 Pig genotype II, 8 cattle, 453 manure treatment, 381 molecular epidemiology genotyping Cryptosporidium in natural water, 149 role of animals in transmission to humans, 138 pigs, 466, 468, 470, 471 sheep, 465 Pig genotypes, neighbor-joining analysis of SSU rRNA gene, 131 Plant-type enzymes, 65, 70 Plasmids, immunization with, 224, 226 Plasmodium, 491, 506 biochemistry, 57, 59, 65 amino acid metabolism, 65 DNA and RNA metabolism, 68 fatty acid metabolism, 62, 63, 64 nucleotide metabolism, 62 structural proteins, 66 genomics comparative, 48
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expressed sequence tags, 44 phylogenetic relationships, 48, 50, 51 sequence projects, 45–46 surface proteins, 48, 50 Polyamine metabolism, 60, 65 Polyclonal antibodies commercially available, 185 from crude antigen, 223–224 from recombinant antigen, 224 Polymerase chain reaction (PCR) cattle, 454 diagnostic methods, 174 inhibitors in environmental samples, 125–126 inhibitors in stool samples, 194–195 mixed infections, 201 nucleic acid detection, 197–203 oocyst negative stool samples, 183 quantitative real-time, 201–202 Taq polymerase inhibitors, 196 typing and subtyping methods, 202 functional genomic analysis, 47 molecular epidemiology, 82 genotyping, 164 genotyping Cryptosporidium in natural water, 148, 149 genotyping tools, 122 human infections, 134 multilocus sequence analysis, 127–128 new genotype characterization, 120, 121 SSU 18S rRNA RFLP, 167 SSU rRNA, sensitivity of nested PCR, 147 waterborne outbreaks, 126 oocyst detection in beverages, 295 oocyst detection in shellfish, 297 pigs, 468 in vitro development studies, 518 water samples, 314 Polymerase chain reaction (PCR)-RFLP diagnostic methods, reporting results, 203 molecular epidemiology, 82 human infections, 134 SSU rRNA gene, 122 SSU rRNA gene, genotyping oocysts in fecal and environmental water samples, 165–167 SSU rRNA gene, water samples, 126 water samples, 126 pigs, 468 Polymerase chain reaction (PCR)-sequencing GP60, genotyping oocysts in fecal and environmental water samples, 167–169 water samples, 126 Polymorphic markers, MLST of C. parvum, C. hominis, and C. meleagridis, 169 Polymorphism, 3′, 125 Poly-T, 134 Polyvinyl alcohol (PVA), 176 Polyvinylpolypyrrolidone (PVP), 195 Population density, factors contributing to transmission in recreational waters, 336 Population genetics
computer programs, 169 molecular epidemiology, 127–130 C. hominis, 129–130 C. parvum, 128–129 MLT and MLST, 127–128 significance of in diagnostics and epidemiology, 130 Population immunity, 85–86 Post-infection sequelae, see Sequelae Postmortem examinations, reptiles, 391 Posttranscriptional gene silencing, lack of, 69 Potassium dichromate, 176, 519 disinfection with, 34 oocyst storage, 164, 176, 186, 195, 407, 506 stool specimen processing, 507, 508, 509, 510 Praziquantel, 258 Prednisolone, latent infection, 151 Prepatent period, 19, 80, 145, 242 birds, 397, 403, 406 infectivity trials, 242 life cycle, 19–20 pigs, 467 Preservation, oocyst, 176; see also Potassium dichromate Pressure, and oocyst viability, 30 Prevalence cattle, 455–458, 459–460, 461, 462 age of animal and, 461, 462 beef, 457, 458 dairy, 456, 458 endemic human cryptosporidiosis, 139–140 geographic distribution, 95 goats, 466 pigs, 468 age and, 468, 470 microscopic detection, 469–470 seroepidemiology, 85 Prevention, control, and treatment, 3, 28–35 animal cryptosporidiosis, 275, 279 avian cryptosporidiosis, 399 chemicals reducing oocyst viability, 31–33, 34–35 drug treatment, see Chemotherapy and prophylaxis environmental factors reducing oocyst numbers, 33, 35 environment disinfection animal models for testing, 31, 485 chemical, 31–33, 34–35 physical, 29–30 epidemiology, 93, 97 factors contributing to transmission in recreational waters, 336 physical factors reducing oocyst viability, 28–31 reptiles, 391 swimming pools, 338 waste management and, 382; see also Waste management waterborne outbreaks, see also Treatment, water drinking water, 325 recreational water contamination, 359, 360–361 wildlife care, 427
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Index zoo animals, 426, 427 Previous infection, see Prior infection Prezwalski’s wild horse, 444 Primates, 424–426 host species, 5–6 laboratory animals, 427 molecular epidemiology, 132 zoo, 426 Primers, PCR functional genomic analysis, 47 mixed infections, minor species detection, 123 multilocus sequence analysis, 170–171 new genotype characterization, 120, 121 and specificity of PCR, 122 Prior infection epidemiology human volunteer studies, 98 incubation period, 81 seroepidemiology, 85–86 Prokaryotic genes/gene products drug targets, 70 horizontal (interspecies) gene transfer, 49, 51 in nuclear genome, 46 Prophylaxis, see Chemotherapy and prophylaxis Protease inhibitors, 263, 264, 278 Proteins, 45 crystal structures, 47 functional genomic analysis, 47 recombinant expression systems, 70–71 surface, 49, 50 Protein Structure Database (PDB), 48 Proteomics, 4, 47 Provamincina® (spiramycin), 257 Public education, recreational water use, 358 Pulsed light, and oocyst viability, 30 Purification of oocysts, see Oocysts Purine biosynthesis, absence of, 51 Pyrimidine biosynthesis, absence of, 51, 62
Q QIAamp kits, 164, 165, 195, 295, 296 Quality, oocyst, 98 Quality assurance oocyst and oocyst DNA shipping for, 203 water samples, 314 Quantification of developing parasites in culture, 516–519 Quantitative real-time polymerase chain reaction (PCR), 201–202
R Rabbit genotype, 8, 167 Rabbits, 3, 423–424 Raccoon genotype, 8 Raccoons, 132, 422 Radiation, and oocyst viability, 29, 30, 225, 299 Rainfall
553 factors contributing to transmission in recreational waters, 336 and manure-treated soil, 380 recreational water contamination, 357 and waterborne outbreaks, 90, 91, 92, 95, 96 Random amplified polymorphic DNA polymerase chain reaction (RAPD-PCR), 122 RANTES, 211, 242 Rats, 422 chemotherapy and prophylaxis, 269–273 C. muris experimental transmission to, 421 host species, 6 Real-time polymerase chain reaction (PCR) genotyping Cryptosporidium in natural water, 149 source tracking, 147 water samples, 126, 314 Recombinant proteins enzyme-linked immunosorbent assay, 192–193 polyclonal antibodies from immunization with antigens, 224 seroepidemiology, 85 Recombinant thrombospondin-related adhesive protein-1, 85, 98 Recombination expression systems for cryptosporidial proteins, 70–71 interspecific, 69 Recombination Detecting Program (RDP), 169 Recreational water, 335–361; see also Waterborne disease case surveillance and epidemiologic studies of associated risk factors, 339–341 endemic human cryptosporidiosis, 140 epidemiology, 92–93 sporadic disease, 97 travel, 94 factors contributing to transmission via, 336–339 outbreaks, 341–357 countries other than United States, 345–347, 350 United States, 341–344, 347–350 outbreaks by water body types and associated deficiencies, 350–357 transmission routes, 3 types and associated deficiencies disinfected water, 352–357 freshwater lakes, rivers, and hot springs, 351–352 marine water, 350–351 types and prevention, 357–360 disinfected water, 358–360 marine and fresh water, 357–358 Regulation, gene, 48 Regulation, transcription, 69 Regulations/regulatory requirements drinking water, 316–318 United Kingdom, 316–317 United States, 317–318 EPA Surface Water Treatment Rule (SWTR), 306 food and drink, 299 Information Collection Rule (ICR), 308, 314, 315
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swimming pools, 338 water monitoring methods, 314 water samples, 150 Regurgitated material, diagnosis in reptiles, 390 Reinfection, epidemiology, 83 Renal cryptosporidiosis, C. baileyi, 406 Repeats, trinucleotide, 123, 124, 125 Replication protein A, 69 Reporting, PCR/RFLP/sequencing results, 203 Reptiles, 3, 389–392 chemotherapy and prophylaxis, 273, 276 clinical signs and pathology, 389–390 diagnosis, 390–391 host specificity, 10, 11 prevention and control, 391 species and genotypes, 130–132 transmission studies, 392 transmission to humans, 138 treatment, 391 Research animals, 487–492 cryptosporidiosis in, 426–427 models of cryptosporidiosis and treatment, 485–493; see also Animal models Reserves, nature, 420 Reservoirs, wildlife species, 420 Respiratory disease, 11, 240 birds, 398 C. baileyi, 396, 397, 405–406 clinical manifestations, 256 life cycle, 11 pigs, 468 Restriction fragment length polymorphism (RFLP), 82, 164, 196 diagnostic methods, 174, 199–200 gene copies and, 122 genotyping tools, 122 pigs, 468 reporting results, 203 SSU rRNA gene, 166–167 typing and subtyping methods, 202 Retroviral therapy, see HAART therapy Reverse transcriptase inhibitors, non-nucleoside (NNRTIs), 263 Reverse transcriptase polymerase chain reaction (RTPCR) genotyping Cryptosporidium in natural water, 148 genotyping tools, 122 subtyping tools, 125 in vitro development studies, 518 water samples, 314 Rhesus monkey, SIV-infected, 488, 490 Ribosomal RNA, 46; see also SSU 18S rRNA gene Rifabutin (Mycobutin®), 258, 261 Risk assessment, 99 Risk factors, 87, 88–89, 97 Ritonavir, 260 Rivers, 351–352 RNA extrachromosomal double stranded, 202 from intracellular developmental stages, 47
ribosomal small subunit gene, see SSU 18S rRNA gene small double-stranded, 82 RNA-mediated posttranscriptional gene silencing, lack of, 69 RNA metabolism, 58–59 RNA splicing, 69 Rodents, 422–423 animal models, 485, 488–489, 490 aquatic, 429 chemotherapy and prophylaxis in, 265, 266–273 host species, 6 transmission routes in animals, 132 wild, 428 Round robin testing, 203 Rovamycina® Rovamycin®, see Spiramycin Roxithromycin®, see Clarithromycin rRNA genes, 46; see also SSU 18S rRNA gene Ruminants, 420–421 chemotherapy and prophylaxis in, 265, 273, 274–275 molecular epidemiology endemic human cryptosporidiosis, 139–140 role of animals in transmission to humans, 138 transmission routes, 132–133 pathogenic species, 175 zoo, 426 Runoff control, 381 Rural areas endemic human cryptosporidiosis, 139–140 waterborne outbreaks, 92
S Safranin methylene blue, 186 Salinity, oocyst inactivation, 298 Salt solution, oocyst flotation method, 182, 183, 509 Salvage pathways, nucleotide, 46, 51, 62 Sample collection stool specimens, 507 water samples, 309 Sand filtration, wastewater effluent, 372, 375 Sandostatin, 264 Sandwich ELISA, 190 Sanitary surveys, 357 Sanitation/hygiene, see Prevention, control, and treatment Saquinavir, 259, 260 Sarcocystis oocyst structure, 21 sequence projects, 45–46 sporogony, 19 taxonomy, 4, 6, 8, 10 Satellite analysis, see Minisatellite/microsatellite analysis Schizogony (merogony), 16, 18, 19, 20 Seal genotypes, 8 Seasonality epidemiology, 95–96 endemic human cryptosporidiosis, 140
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Index and intervention, 99, 100 source tracking, 146–147 recreational water contamination, 339 wild animal oocyte shedding in California, 420 zoo animal oocyst shedding, 426 Secondary PCR product detection, 168 Secretory IgA, 222–223 Sedimentation fecal samples, 487, 502 oocysts adherence to soil particles and, 380 purification of, 194 in water, 297, 298, 308 sporozoites, 502 water treatment, 308, 318, 319, 376 Semi-nested PCR, 149 Sensitivity of detection cattle, 454 diagnostic methods, 190–191 molecular epidemiology, 82 PCR nested, 183 RT-PCR format, 125 Sequelae C. parvum versus C. hominis, 146 duration of symptoms, 81–82 in humans, 175 infectivity trials, 244 Sequence analysis, genome survey sequences, 44–45 Sequence differences, copy number and, 7, 125 Sequence length polymorphisms, C. hominis population genetics, 129 Sequences genome survey, 44–45 genomic, 45 Sequencing, 197 C. bovis, 200 databases, 52–53 diagnostic methods, 174, 197 C. bovis, 200 reporting results, 203 typing and subtyping methods, 202 genotyping tools, 122 molecular epidemiology, 82 genotyping, 122, 164 GP60, 168–169 multilocus sequence analysis, 127–128 subtyping tools, 123 water samples, 126 multilocus sequence analysis, 127–128 pigs, 468 typing and subtyping methods, 202 Serial passage and virulence, 453–454 Seroconversion, population immunity, 85 Serodiagnostic assays cattle, 454 ELISA-based, 192–193 Seroepidemiology/population immunity dogs, 441 ELISA-based serodiagnostic assay, 193 epidemiology, 85–86
555 Serum antibody tests, see Antibodies 17kD antigen ELISA-based serodiagnostic assay, 193 enzyme-linked immunoelectrotransfer blot (EITB), 191, 192 seroepidemiology, 85 Severity of infection, 1 coinfection and in fish, 388 in pigs, 467 differences among species infecting humans and agricultural animals, 145, 453 gnotobiotic pig model, 489, 490 host immune status and, 1–2 immune response and, 215, 223, 256 pathophysiology, 241 serial passage and, 453–454 Sewage treatment, 371, 372 Sexual reproduction (gamogony), 16–17, 21, 22, 23 Sexual stages, structure, 21, 22, 23, 25–26 Sheep, 3, 224, 462, 464, 465 chemotherapy and prophylaxis, 275 C. parvum in small mammals and, 83 host species, 5 molecular epidemiology role of animals in transmission to humans, 138 subtype families, 84 transmission routes in animals, 132–133 species infecting, 11 Sheep novel genotype, 8, 11, 465 Shelf life of purified oocysts, 511 Shellfish, 35, 290–291, 295–297; see also Foodborne disease Shipping, oocysts and oocyst DNA, 183, 203 Shrews, 424 Sieving, sporozoite purification, 510 Sieving, stool, 508 Single nucleotide polymorphisms (SNP), 129 Single-strand conformation polymorphism (SSCP) method, 122–123, 125 Site of infection, 11, 178 SIV-infected rhesus monkey model, 488, 490 Sixty kilodalton glycoprotein, see GP60 Size, genome, 45 Skunk genotype, 8 epidemiology, 83 evolutionary relationships of avian isolates, 404 genotyping Cryptosporidium in natural water, 149 Sludge treatment, waste management, 376–378 Small double-stranded RNA, 82 Small subunit ribosomal RNA, see SSU 18S rRNA gene Smear, oocyst, 177 Snake genotype, 8, 167 evolutionary relationships of avian isolates, 404 hosts infected by, 11 Snakes, 3, 273, 276 Sodium acetate-acetic acid formalin (SAF), 176 Sodium chloride, oocyst flotation on brine, 182, 183 Sodium docecyl sulfate, 183, 196, 312
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Sodium hypochlorite, 399, 503, 505, 510, 511, 512, 514, 518 Software oocyst detection, laser scanning cytometry, 313–314 population genetics, 169 Soil freeze-thaw cycles and oocyst viability, 28 oocysts in dairy cattle manure applied to, 380–381 sludge application, 377 Solar irradiation, and oocyst viability, 30 Somatostatin analogs, 264–265 Sources, disease, see also Transmission epidemiology, 87, 97 foodborne Cryptosporidia, 297–298 life cycle, environmental stage, 12 molecular diagnostic methods, PCR, 202–203 molecular epidemiology, 82, 84, 146–151 epidemic human cryptosporidiosis, 144–145 population genetics, significance of, 130 utility of tools, 127 Sources, oocyst, 505–506 Spatial tracking, 130 Species (of Cryptosporidia), 1 and clinical course incubation period, 81 prepatent and patent period, 18 human pathogens, 175 livestock, 175 molecular analysis, 455 pigs, 471 livestock, cattle, 452, 453, 454–455, 457 age of animal and, 461, 462 beef cattle, molecular methods, 460 dairy cattle, molecular methods, 459–460 mixed dairy and beef cattle, molecular methods, 461 molecular epidemiology, 150 biological, clinical, and epidemiologic differences among, 145–146 concepts for species, genotypes, and subtypes, 120–121 host range, 84 human cryptosporidiosis, 133–138 intraspecific population structure, 127 in mammals, birds, reptiles, and fish, 130–132 methods for identification of, 120–121 typing and subtyping methods, 202 utility of tools, 126 nomenclature, genotype versus, 84–85 number of, 2, 174–175 waterborne outbreaks, 126, 150 wild mammals, 428–429 Species distributions, 84 endemic human cryptosporidiosis, 139 oocysts, 126 Species divergence, 127 Specificity of molecular techniques, see also specific methods conserved sequences and, 122
interspecies differences versus intragenotypic variation, 7 Spectroscopy, water samples, 314 Spectrum of infectivity, 83 Spermine-spermidine, 65 Spider monkeys, 425 Spiramycin, 257, 260 Splash pads, 355–356 Sporadic disease, 235 in cattle, 453 Cryptosporidium suis in humans, 470 epidemiology, 84, 96, 97 C. parvum versus C.hominis, 84, 144, 146 intervention effects, 99 person-to-person transmission, 93 risk factors, 87 role of animals in transmission to humans, 138 typing/subtyping/subtypes, 82, 144 mixed infections and, 201 recreational water-associated, 340, 351 Sporadic disease, epidemiology, 87, 97, 340 Sporogony, 18–19, 24 Sporozoites attachment for feeder organelle, 23 chorioallantoic membrane culture, 520 DAPI staining, 189 expressed sequence tags, 44 glycoproteins, 67 life cycle, see Life cycle morphology, see Morphology purification of, 513 structure, 17, 18, 22–23 taxonomic classification of cryptosporidia, 4 in vitro systems, see In vitro systems/cell culture Spray features, 355–356 Squirrel(s), 11, 423, 429 Squirrel genotype, 11, 149 Squirrel monkeys, 424 SspI, 164, 166, 167, 199, 200 SSU 18S rRNA gene avian Cryptosporidium species, 411, 412 C. galli, 410 evolutionary relationships, 404 genetic similarities, 407 cattle, 454–455 species and genotypes, 461 species and genotypes in dairy cattle, 459, 460 species and genotypes infecting beef cattle, 460 C. meleagridis, 403 diagnostic methods, 197–200 mixed infections, 201 PCR, primers and step cycle protocols, 197, 198 typing and subtyping methods, 202 molecular diagnostic methods, 197–200 molecular epidemiology animal cryptosporidiosis, 131 C. canis heterogeneity, 132 C. parvum population genetics, 129 Cryptosporidium taxonomy, 120 genotyping, 164
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Index genotyping Cryptosporidium in natural water, 148, 149 genotyping oocysts in fecal and environmental water samples, 165–167 genotyping tools, 122 human infections, 134 nested PCR, surface water, 147 PCR primers, 121 water samples, 126 oocyst detection in shellfish, 296 phylogenetic analysis, 50 pigs, 470, 471 sheep, 465 species identification, 7 Staining biopsy, 193 developing parasites in culture, 516–517 oocysts, 179, 186–188 auramine phenol, 186–187 cattle, 454 commercial diagnostic kits, 180–181 comparison of diagnostic methods, 191 horse samples, 443 Ziehl-Neelsen stain, modified, 187–188 Stavudine, 259, 260 Stool specimens, 176–178, 179 collection, 507 diagnosis in reptiles, 390 diagnostic tests, see Diagnostic methods oocyst genotyping and subtyping, 164–169 DNA extraction from fecal samples using QIAamp kit, 164 DNA extraction from water and fecal samples using FastDNA SPIN Soil kit, 165 genotyping by PCR/RFLP analysis of SSU rRNA gene, 165–167 subtyping by PCR sequencing analysis of GP60 gene, 167–169 oocyst negative, 183 oocyst purification, 507–510 PCR inhibitors, 194–195 preservatives, 176 stool specimen preservation, 176 Storage oocysts, 506 and infectivity, 511 and viability, 30 stool specimens, 176 Storm water genotypes, 373 Strains epidemiology, 83, 87, 97 nomenclature, 85 Structural genomics, 47 Structural proteins, 66–67 Structure, 21–26, 27 attachment for feeder organelle, 14, 15, 23–24 and life cycle, 4, 6 oocyst, 21–22, 24, 25, 26 sexual stages, 21, 22, 23, 25–26 sporozoites, 17, 18, 22–23 taxonomic classification of cryptosporidia, 4
557 trophozoites and merozoites, 17, 19, 24–25, 27 Subclinical infections, see Asymptomatic/subclinical infection Subgenotypes, 7–8, 121 Subgroups, C. parvum isolates, 83 Substance P, 241 Subtypes C. meleagridis, 403 C. parvum, cattle, 463–464 epidemiology, 83 IgG antibody response to reference strain of C. parvum, 85–86 sources, 87 molecular epidemiology, 84, 150 animal transmission routes, 133 C. hominis population genetics, 129 concepts, 120–121 endemic human cryptosporidiosis, 140, 141 epidemic human cryptosporidiosis, 142–143, 144–145 linkage disequilibrium, 128 occurrence in humans versus sheep and cattle, 83 nomenclature, 85 Subtyping, 3–4 cattle, 458, 460–461, 463–464 molecular epidemiology, 82, 123–125 epidemic human cryptosporidiosis, 144–145 oocysts in fecal and environmental water samples, 164–169 PCR, 197, 202–203 Sucrose flotation method, 312, 500, 509; see also Flotation, oocysts Sucrose flotation method, oocyst concentration, 178, 182–183 Sucrose gradients, 509, 513 Sulfaquinoxaline, 265 Supportive therapy Surface proteins, 7, 49, 50, 67–68 Surface water, see Waterborne disease Surveillance, epidemiology, 80 Swimming, 142–143, 144, 145, 340; see also Recreational water Swimming pools, 352–355 operation practices, 359 regulation of, 338 Symptoms, see Clinical disease and pathology
T Taq polymerase inhibitors, 196 Tasmanian pademelon, 424 TATA-binding protein, 69 Taurocholate treatment, oocyst, 514 Taxonomy, 4–10, 120 C. parvum nomenclature, 84 genotypes, 8 historical developments, 3 interspecies differences versus intragenotypic variation, 7
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mammalian host species, 5–6 molecular methods, 3–4 naming caveats, 121 TCA, TCG, TCT repeats, 123, 124, 125, 463–464 T-cell mediated immunity, 213–217, 225; see also Cellmediated immune response HAART therapy and, 263 importance of T cells in infection control, 213–215 mucosal compartments, 215–216 parasite antigen-specific priming of T-cells and Tcell memory, 216–217 Temperature chorioallantoic membrane culture, 520 dairy cattle manure treatment, 379–380 and oocyst infectivity, 511 oocyst sensitivity, 98, 298, 299 and oocyst viability, 28, 29, 30 sludge treatment, 377 and water disinfection, 321–322 zoo animals, 426 Temporal factors descriptive and analytical investigations, 97 molecular epidemiology population genetics, 127 population genetics, significance of, 130 utility of tools, 126 Terminology, genotypes and subtypes, 121 Th1 cytokines, 217–219 Th2 cytokines, 219–220, 225 Thrombospondin-related adhesive proteins (TRAPs), 68, 85, 98 Thymidine kinase (TK), 51, 52, 62 Time course of infections, see also Clinical disease and pathology cattle, 452 enzyme-linked immunoelectrotransfer blot (EITB), 191–192 in humans, 175 antibody appearance, 85 clinical manifestations, 256 elderly, 238 versus gnotobiotic pig model, 146 infectivity trials, 242 pigs, 467 prepatent and patent period, 19–20 Toddler pool, 340 Toll-like receptors, immune system, 211 Tortoise genotype, 8, 167 Toxoplasma, 21 biochemistry, 57, 59, 62, 63, 64 chemotherapy, 257, 263 coding capacity, 46 expressed sequence tags, 44 gene regulation, 48 genome properties, 45 in mollusks, 291 oocyst structure, 19 sequence projects, 45–46 taxonomy, 4, 6 tritiated uracil incorporation, 518
Tracking of infection, population genetics and, 130 Trade animals, 427 Transmission, 3, 120; see also Sources, disease amphibial cryptosporidiosis, 389 biology, 26, 28 early studies, 9 epidemiology, 92, 93 descriptive and analytical investigations, 97 drinking water, 90–92 food and drink, 93–94 other drinking waters, 92 person-to-person, 93 recreational waters, 92–93 travel, 94 zoonotic, 93 fish cryptosporidiosis, 388 livestock, 452 molecular epidemiology, 84, 150 animal cryptosporidiosis, 132–133 C. parvum population genetics, 129 endemic human cryptosporidiosis, 139–141 genotype and subtype identification, 121 population genetics, significance of, 130 utility of tools, 127 reptilian cryptosporidiosis, 392 wild mammals, 428 Transporters, 46, 67–68 genomics, 52 highly streamlined metabolism, 58–59 TRAP C1, 197 TRAP C2, 148 Travel epidemiology, 89, 95, 97 recreational water use caveats, 358 Treatment, clinical, see also Chemotherapy and prophylaxis latent infection, 151 reptiles, 391 Treatment, environment, see Prevention, control, and treatment Treatment, water, 318–325 animal models for testing, 492 disinfection, 320–325 filtration, 318–320 and outbreak incidence, 348, 349, 350 Trinucleotide repeats, GP60, 123, 124, 125, 463–464 tRNA, 46, 70 Trophozoites, 17, 19, 24–25, 27, 513 Tubulins, 66, 69, 70, 202 Tumor necrosis factor-alpha, 220–221, 241 Turbidity and immunomagnetic separation, 184 water, and waterborne outbreaks, 92 Turkeys, 3, 11 27kD antigen, 85, 191, 192, 193 Typing molecular epidemiology, 82, 84 nomenclature, 84–85 PCR, 202–203 waterborne outbreaks, 344
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Index
U Ultrasound, 29, 299 Ultraviolet light DNA repair mechanisms, 68–69 oocyst sensitivity, 299 and oocyst viability, 29, 30 wastewater effluent treatment, 372, 374 water disinfection, 322–325 Uncommon species, C. parvum antibody reactivity, 190 United States EPA Surface Water Treatment Rule (SWTR), 306 recreational water outbreaks, 341–344, 347–350 Urban areas endemic human cryptosporidiosis, 139–140 source tracking, 147 Uridine kinase-uracilphosphoribotransferase (UKUPRT), 51, 62
V Vaccination, 223–225 Vapreotide, 264–265 Vegetables, 290, 293–294 Vinblastine, 277 Viral therapy, 255; see also HAART therapy Virulence, 256 animal models, 486, 492 attenuation for vaccines, 225 C. meleagridis, 402 C. parvum in calves, 453, 454 human volunteer studies, 98 molecular epidemiology, 150 serial passage through calves versus mice, and attenuation, 453, 454 variations in, 7–8 Virus coinfections birds, 397, 398 pigs, 467 Viruslike double-stranded RNA, 69; see also Doublestranded RNA Vortex flow filtration, 312 VspI, 164, 166, 167, 199, 200
W Waste dumping, 357 Waste management, 371–382 beef cattle manure and treatment, 381 biological treatment, 376 clarifiers, 375 dairy cattle manure and treatment, 378–381 effluent treatment, 372–375 overall effects of treatments, 376 sludge treatment, 376–378 swine manure and treatment, 381–382 Wastewater effluent treatment, 372–375 chlorine dioxide, 374 general, 373–374
559 ozonation, 374 sand filtration, 375 UV light, 374 factors contributing to transmission in recreational waters, 336 genotyping Cryptosporidium in natural water, 148, 149 source tracking, 147 Waterborne disease, 3; see also Recreational water environmental factors reducing oocyst numbers, 33 epidemiology, 88 burden of disease, 96 incubation period, 80, 81 intervention, 99 protection of vulnerable populations, 95 risk assessment, 99 seasonality, 95–96 sporadic disease, 97 surveillance, 80 transmission routes, C. hominis, 90 transmission routes, C. parvum, 89 travel, 94 food contamination, 289, 290, 297–298 historical developments, 3 molecular epidemiology, 84 endemic human cryptosporidiosis, 140 epidemic human cryptosporidiosis, 141, 142–143, 144–145 source tracking, 146–147 transmission routes in animals, 132 species and genotypes of oocysts, 126 transmission routes, 3, 120 watershed contamination, domestic animals and wildlife as sources, 429–430 Waterborne Disease and Outbreak Surveillance System (WBDOSS), 341, 344 Water buffalo, 5, 471 Water parks, 338, 352–355 Water potential, and oocyst viability, 28, 29 Water samples DNA extraction from, 165 genotyping by PCR/RFLP analysis of SSU rRNA gene, 165–167 subtyping by PCR sequencing analysis of GP60 gene, 167–169 Watershed land, recreational water contamination, 357 Weather factors contributing to transmission in recreational waters, 336 and waterborne outbreaks, 89, 90, 91, 92, 95, 96 Western blot, 454 Wild animals, 420–426 bats, 424 carnivores, 421–422 insectivores, 424 lagomorphs, 423–424 marsupials, 424 nonhuman primates, 424–426 omnivores, 421
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rodents, 422–423 ruminants, 420–421 transmission routes, 3, 427–429 C. parvum, 89 domestic animal cross transmission, 419 farm animal cross transmission, 420 role in transmission to humans, 138 as source of infection in surface water, 147 transmission routes in animals, 132 transmission to humans, 138 Wildlife genotypes source tracking, 146–147 wastewater effluent oocysts, 373 Woodcock genotype, 8
X Xiao et al. assay, 121, 198–199
XR2/XF2, 199
Z Zalcitabine, 259, 260 Zidovudine, 259 Ziehl-Neelsen stain, modified, 174, 177, 182, 186, 187–188, 191 Zirconia bead extraction, 194 Zithromax®, see Azithromycin Zoo animals, 26, 138, 421, 426–427, 428, 429 Zoonotic transmission, see Animal transmission/zoonotic disease Zoos, recreational water contamination, 355–356 Zygote, 24
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FIGURE 2.2 Domain organizations of a representative set of surface proteins from Cryptosporidium parvum (top panel) and orthologs common to Plasmodium falciparum (bottom panel). All proteins shown here have a signal peptide sequence represented by a rectangular box at the beginning of the architectures. The domains are labeled as in Templeton et al., 2004a. (Reproduced with permission from Genome Research; Templeton, T.J., Iyer, L.M., Anantharaman, V., Enomoto, S., Abrahante, J.E., Subramanian, G.M., Hoffman, S.L., Abrahamsen, M.S., and Aravind, L. 2004a. Comparative analysis of apicomplexa and genomic diversity in eukaryotes. Genome Res. 14, 1686–1695.)
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FIGURE 3.1 Major metabolic pathways and their connections in Cryptosporidium based on genome annotation and recent biochemical data. Solid arrow bars indicate direct biochemical reactions, whereas dashed lines indicate multiple-step reactions or connections between pathways. Most enzymes are highlighted with solid dark green boxes, but those involved in ATP consumption or production are highlighted with orange and red boxes, respectively. Organic end products are boxed. Abbreviations: ABC, ATPbinding cassette transporter; ACBP, acyl-CoA binding protein; ACC, acetyl-CoA carboxylase; AceCL, acetate-CoA ligase (also termed acetyl-CoA synthetase); ACL, fatty acid-CoA ligase (also termed fatty acyl-CoA synthetase); ADC, arginine decarboxylase; ADH, alcohol dehydrogenase; adhE type E alcohol dehydrogenase; AIH, agmatine iminohydrolase; AK, adenosine kinase; AMPDA, AMP deaminase; dCMPDA, dCMP deaminase; CTPS, CTP synthase; DHF, dihydrofolate; DHFR, dihydrofolate reductase; CMPK, CMP kinase; CMPS, CMP synthase; GAPDH, glyceraldehyde phosphate dehydrogenase; GDH, glycerol phosphate dehydrogenase; GGH, glucoside glucohydrolase; GS, glutamine synthetase; HK, hexokinase; IMPDH, IMP dehydrogenase; LCE, long-chain fatty acyl elongase; LDH, lactate dehydrogenase; MDH, malate dehydrogenase; ME, malic enzyme; MPGT, mannose-1-phosphate guanylyltransferase; mTHF, methylene tetrahydrofolate; ORP, oxysterol-binding protein-related protein; PDC, pyruvate decarboxylase; PEPCL, phosphoenolpyruvate carboxylase; PFK, phosphofructokinase; PGI, phosphoglucose isomerase; PGK, phosphoglycerate kinase; PGM, phosphoglycerate mutase; PGluM, phosphoglucose mutase; PK, pyruvate kinase; PKS, polyketide synthase; PMI, mannose-6-phosphate isomerase; PMM, phosphomannomutase; PNO, pyruvate NADP+ oxidoreductase; PV, parasitophorous vacuole; PVM, parasitophorous vacuole membrane; RNR, ribonucleoside-diphosphate reductase; SHMT, serine hydroxymethyl transferase; SpdSyn, spermidine synthase; SpmSyn, spermine synthase; SSAT, spermidine:spermine N1-acetyltransferase; THF, tetrahydrofolate; TIM, triosephosphate isomerase; TIM17, translocase of inner mitochondrial membrane (17 kDa); TK, thymidine kinase; TOM40, translocase of outer mitochondrial membrane (40 kDa); TP, trehalose phosphatase; TPS, trehalose phosphate synthase; TS, thymidylate synthase; UDPGPP, UDP-glucose pyrophosphorylase; UDPK, UDP/CDP kinase; UK, uridine kinase; UMPK, UMP/CMP kinase; UPRT, uracil phosphoribosyltransferase.
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A
B
C
D
FIGURE 6.1 Panel of oocyst images. (A) Two C. parvum oocysts following cesium chloride purification and photographed in suspension using Nomarski differential interference contrast microscopy. Note the presence of three of the four sporozoites in the upper oocyst and the typical padlock arrangement of sporozoites in the lower oocyst. Magnification 1250×. (B) A group of C. parvum oocysts prepared from an air-dried formol ether concentrate of stool and stained with the modified Ziehl–Neelsen acid-fast stain. Cryptosporidium oocysts stain red and appear as spherules. Note the variation in staining intensity of individual oocysts in this image. The contents of many oocysts will appear amorphous, but some contain the characteristic crescentic forms of sporozoites. Magnification 1250×. (C) A group of C. parvum oocysts prepared from an air-dried formol ether concentrate of stool and stained with the auramine phenol stain. Cryptosporidium oocysts appear ring shaped and exhibit a characteristically bright green fluorescence against a dark background. Magnification 1250×. (D) A group of C. parvum oocysts prepared from an air-dried formol ether concentrate of stool and stained with a commercially available fluorescein isothiocyanate-labeled monoclonal antibody reactive with exposed epitopes on Cryptosporidium spp. oocysts. Note the following three characteristics of Cryptosporidium spp. oocysts using this stain: (a) characteristic apple-green fluorescence delineating the oocyst wall, under the FITC filter set. Often the fluorescence has an increased intensity around the entire circumference of the oocyst, with no visible breaks in oocyst wall staining; (b) round or slightly ovoid objects; (c) a size of 4 to 6 µm in diameter for most human pathogens. Magnification 1250×.
Type of Exposure (n=189)
Etiologic Agent (n=189) Norovirus 11.1%
Treated water 49.7%
Cryptosporidium spp. 33.3%
Shigella spp. 15.9% E. coli 10.6%
Untreated water 50.3%
Giardia spp. 7.4%
Otherb 4.2%
Unidentified 17.5%
Etiologic Agent: Untreated Water (n=95) Shigella spp. 24.2%
Otherb 3.2%
E. coli 16.8%
Cryptosporidium spp. 59.6%
Norovirus 13.7% Otherb 5.3% Cryptosporidium spp. 7.4%
Unidentified 27.4%
Etiologic Agent: Treated Water (n=94)
Giardia spp. 7.4%
Unidentified 7.4% Shigella sonnei 7.4% Giardia intestinalis 7.4%
Norovirus 8.5% E. coli 4.3%
FIGURE 12.6 Recreational water-associated outbreaks of acute gastroenteritis by type of exposure and etiologic agent—United States, 1988–2004.a aFive outbreaks involved Cryptosporidium with one or more other pathogens detected in greater numbers; these outbreaks have been classified as non-Cryptosporidium outbreaks; bThese include outbreaks of Campylobacter, Plesiomonas, Salmonella, and suspected chemical or toxin etiologies.
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Type of Exposure (n=20,441)
Etiologic Agent (n=20,441)
Treated water 73.7%
Cryptosporidium spp. 70.7%
Unidentified/ Otherb 11.2% Giardia spp. 2.3%
Untreated water 26.3%
E. coli 2.3% Shigella spp. 9.4%
Etiologic Agent: Untreated Water (n=5,386) Shigella spp. 31.5%
Norovirus 4.3%
Etiologic Agent: Treated Water (n=15,055)
E. coli 7.7%
Cryptosporidium spp. 91.6%
Norovirus 9.5%
Unidentified/ Otherb 1.7% Shigella sonnei 1.5% Cryptosporidium Giardia intestinalis 2.5% spp. 12.2% E. coli 0.4% Unidentified/ Otherb 37.2%
Giardia spp. 1.7%
Norovirus 2.4%
FIGURE 12.7 Recreational water-associated cases of acute gastroenteritis by type of exposure and etiologic agent—United States, 1988–2004a. aFive outbreaks involved Cryptosporidium with one or more other pathogens detected in greater numbers; the cases attributed to these outbreaks have been classified as non-Cryptosporidium cases; bOther includes outbreaks of Campylobacter, Plesiomonas, Salmonella, Cyanobacter, and suspected chemical etiologies.
FIGURE 20.3 Panels A, B, and C illustrate the progressive gradient techniques used to purify C. parvum oocysts. Panel A shows primary (left) and secondary (right) discontinuous sucrose gradients after centrifugation with the light debris (d) at the interface between the sample and the STP 1:4 layer, the oocysts (o) at the interface between the STP 1:4 and STP 1:2 layers, and the pellet (p) of dense debris. Panel B shows examples of the cesium chloride step gradients: the first tube (left) shows the sucrose gradientpurified oocyst sample (s) layered over the cesium chloride column before centrifugation; the second tube (middle) shows the same tube after centrifugation with the oocysts (o) at the interface between the sample and the cesium chloride and a small pellet (p) of dense debris at the bottom; the third tube (right) shows the purified pellet of oocysts (o) after the saline wash step. Panel C shows an example of a Percoll® gradient used to isolate sporozoites (sp) and empty or partially empty oocyst walls (w). Panel D shows a differential interference contrast (DIC) view of representative C. parvum oocysts purified using the sucrose and cesium chloride gradient techniques. Panel E shows purified sporozoites labeled with a monoclonal antibody specific for a sporozoite surface protein (C3B4, reacts with the 23-kDa surface antigen); this view includes a secondary antibody conjugated with fluorescein (green). Panel F shows the same antibody labeling of merozoites emerging from a meront in a paraffin section of infected mouse ileum. Panels G and H show, respectively, the DIC view and immunofluorescent view (C3B4 labeling) of budding merozoites in a cell culture preparation. Panels I, J, and K show, in sequence, the DIC view of a late trophozoite/early meront, DAPI nuclear staining of the host cell and the parasite nuclei (above host nucleus, see arrow) and, finally, the intracellular label of the parasite by the panreactive monoclonal antibody C3C3 (conjugated with the red fluorescent dye Cy3). Panel L shows C. parvum growth in MDCK cells at 24 h post inoculation (PI) where the oocyst (green) has been labeled directly with FTSC and the developing stages (mostly early trophozoites/Type II meronts) have been labeled with the monoclonal antibody C3C-Cy3. Panel M shows MDCK cells at 48 h PI; note the obvious FTSC-labeled oocyst and the distinct C3C3-Cy3 labeling of the gamonts and meronts. Scale bar applies to panels D through M.
