Cryptosporidium and Cryptosporidiosis

52268_C000.fm Page i Monday, October 15, 2007 6:44 AM CRYPTOSPORIDIUM AND CRYPTOSPORIDIOSIS Second Edition 52268_C0...

Author: Ronald Fayer | Lihua Xiao

96 downloads 1999 Views 15MB Size Report

This content was uploaded by our users and we assume good faith they have the permission to share this book. If you own the copyright to this book and it is wrongfully on our website, we offer a simple DMCA procedure to remove your content from our site. Start by pressing the button below!
Report copyright / DMCA form

DOWNLOAD PDF

52268_C000.fm Page i Monday, October 15, 2007 6:44 AM

CRYPTOSPORIDIUM AND

CRYPTOSPORIDIOSIS Second Edition

52268_C000.fm Page ii Monday, October 15, 2007 6:44 AM

52268_C000.fm Page iii Monday, October 15, 2007 6:44 AM

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

52268_C000.fm Page iv Monday, October 15, 2007 6:44 AM

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

52268_C000.fm Page v Monday, October 15, 2007 6:44 AM

Ernest Edward Tyzzer (Courtesy of the Francis A. Countway Library of Medicine, Boston, Massachusetts)

52268_C000.fm Page vi Monday, October 15, 2007 6:44 AM

52268_C000.fm Page vii Monday, October 15, 2007 6:44 AM

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

52268_C000.fm Page viii Monday, October 15, 2007 6:44 AM

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

52268_C000.fm Page ix Monday, October 15, 2007 6:44 AM

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.

52268_C000.fm Page x Monday, October 15, 2007 6:44 AM

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

52268_C000.fm Page xi Monday, October 15, 2007 6:44 AM

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.

52268_C000.fm Page xii Monday, October 15, 2007 6:44 AM

52268_C000.fm Page xiii Monday, October 15, 2007 6:44 AM

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

52268_C000.fm Page xiv Monday, October 15, 2007 6:44 AM

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

52268_C001.fm Page 1 Wednesday, September 26, 2007 1:39 PM

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


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).


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


4

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


General Biology

5

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)


6

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


General Biology

7

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.


8

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


General Biology

9

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.


10


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


General Biology

11

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).


12

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.


General Biology

13

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


14


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


General Biology

15

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


16


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


General Biology

17

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.


18


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).


General Biology

19

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


20


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


General Biology

21

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


22


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


General Biology

23

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


24


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


General Biology

25

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


26


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


General Biology

27

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.)


28


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).


General Biology

29

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


30


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.


General Biology

31

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)


32


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.


General Biology

33

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


72% reduction

Excyst


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).


34


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


General Biology

35

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.


36


Augustin-Bichl, G., Boch, J. and Henkel, G. 1984. Kryptosporidien-Infektionen bei Hund und Katze. Berl. Muench. Tieraerztl. Wochenschr. 97, 179–181. Barbee, S.L., Weber, D.J., Sobsey, M.D. and Rutala, W.A. 1999. Inactivation of Cryptosporidium parvum oocyst infectivity by disinfection and sterilization processes. Gastrointest. Endosc. 49, 605–611. Barker, I.K. and Carbonnel, P.L. 1974. Cryptosporidium agni sp. n. from lambs and Cryptosporidium bovis sp. n. from a calf with observations on the oocyst. Z. Parasitenkd. 44, 289–298. Belosevic, M., Craik, S.A., Stafford, J.L., Neuman, N.F., Kruithof, J. and Smith, D.W. 2001. Studies on the resistance/inactivation of Giardia muris cysts and Cryptosporidium parvum oocysts exposed to mediumpressure ultraviolet radiation. FEMS Microbiol. Lett. 204, 197–203. Betancourt, W.Q. and Rose, J.B. 2005. Microbiological assessment of ambient waters and proposed water sources for restoration of a Florida wetland. J. Wat. Hlth. 32, 89–100. Bird, R.G. 1981. Protozoa and viruses. Human cryptosporidiosis and concomitant viral enteritis, in Parasitological Topics, a presentation volume to PCC Garnham, Canning, E.U., Ed., Special publication no. 1, Society of Protozoologists, 39–47. Blewett, D.A. 1989. Disinfection and oocysts, in Proc. 1st Int. Workshop on Cryptosporidiosis, Angus, K.W. and Blewett, D.A., Eds., September 7–8, 1988, Edinburgh, 107–115. Brownstein, D.G., Strandberg, J.D., Montali, R.J., Bush, M. and Fortner, J. 1977. Cryptosporidium in snakes with hypertrophic gastritis. Vet. Pathol. 14, 606–617. Campbell, A.T., Robertson, L.J. and Smith, H.V. 1993. Effects of preservatives on viability of Cryptosporidium parvum oocysts. Appl. Environ. Microbiol. 59, 4361–4362. Campbell, A.T., Robertson, L.J., Snowball, M.R. and Smith, H.V. 1995. Inactivation of oocysts of Cryptosporidium parvum by ultraviolet irradiation. Wat. Res. 29, 2583–2586. Campbell, I., Tzipori, S., Hutchison, G. and Angus, K.W. 1982. Effect of disinfectants on survival of cryptosporidium oocysts. Vet. Rec. 111, 414 and 415. Carreno, R.A., Martin, D.S. and Barta, J.R. 1999. Cryptosporidium is more closely related to the gregarines than to coccidia as shown by phylogenetic analysis of apicomplexan parasites inferred using small-subunit ribosomal RNA gene sequences. Parasitol. Res. 85, 899–904. Castro-Hermida, J.A., Pors, I., Mendez-Hermida, F., Ares-Mazas, E., and Chartier, C. 2006. Evaluation of two commercial disinfectants on the viability and infectivity of Cryptosporidium parvum oocysts. Vet. J. 171, 340–345. Cevallos, A.M., Zhang, X., Waldor, M.K., Jaison, S., Zhou, X., Tzipori, S., Neutra, M.R. and Ward, H.D. 2000. Molecular cloning and expression of a gene encoding Cryptosporidium parvum glycoproteins gp40 and gp15. Infect. Immun. 68, 4108–4116. Chalmers, R.M., Elwin, K., Reilly, W.J., Irvine, H., Thomas, A.L. and Hunter, P.R. 2002. Cryptosporidium in farmed animals: the detection of a novel isolate in sheep. Int. J. Parasitol. 32, 21–26. Clark, J.J. 1894–95. A study of coccidia met with in mice. J. Micro. Sci. 37, 277–283. Collins, M.V., Flick, G.J., Smith, S.A., Fayer, R., Croonenberghs, R., O’Keefe, S. and Lindsay, D.S. 2005. The effect of high pressure processing on infectivity of Cryptosporidium parvum oocysts recovered from experimentally exposed Eastern oysters (Crassostrea virginica). J. Eukaryot. Microbiol. 52, 500–505. Collins, M.V., Flick, G.J., Smith, S.A., Fayer, R., Rubendall, E. and Lindsay, D.S. 2005a. The effects of ebeam irradiation and micrwave energy on Eastern oysters (Crassostrea virginica) experimentally infected with Cryptosporidium parvum. J. Eukaryot. Microbiol. 52, 484–489. Current, W.L. 1985. Cryptosporidiosis, a protozoologist’s view of an emerging zoonosis. Microecol. Ther. 15, 165–200. Current, W.L. 1988. The biology of Cryptosporidium. ASM News 54, 605–611. Current, W.L. and Reese, N.C. 1986. A comparison of endogenous development of three isolates of Cryptosporidium in suckling mice. J. Protozool. 33, 98–108. Current, W.L., Upton, S.J. and Haynes, T.B. 1986. The life cycle of Cryptosporidium baileyi n. sp. (Apicomplex, Cryptosporidiidae) infecting chickens. J. Protozool. 33, 289–296. Craik, S.A., Weldon, D., Finch, G.R., Bolton, J.R. and Belosevic, M. 2001. Inactivation of Cryptosporidium parvum oocysts using medium- and low-pressure ultraviolet radiation. Wat. Res. 35, 1387–1389. Deng, M.Q. and Cliver, D.O. 1999. Cryptosporidium parvum studies with dairy products. Int. J. Food Microbiol. 46, 113–121. Ditrich, O., Palkovic, L., Sterba, J., Prokopic, J., Loudova, J. and Giboda, M. 1991. The first finding of Cryptosporidium baileyi in man. Parasitol. Res. 77, 44–47.


General Biology

37

Drescher, A.C., Greene, D.M. and Gadgil, A.J. 2001. Cryptosporidium inactivation by low-pressure UV in a water disinfection device. J. Environ. Health 64, 31–35. Dunn, J., Ott, T. and Clark, W. 1995. Pulsed light treatment of food and packaging. Food Tech. 49, 95–98. DuPont, H., Chappell, C., Sterling, C.R., Okhuysen, P.C., Rose, J.B. and Jakobowski, W. 1995. The infectivity of Cryptosporidium parvum in healthy volunteers. N. Engl. J. Med. 332, 855–859. Duszynski, D.W. 1969. Two new coccidia (Protozoa, Dimeriidae) from Costa Rican lizards with a review of the Eimeria from lizards. J. Protozool. 16, 581–585. Enemark, H.L., Ahrens, P., Bille-Hansen, V., Heegaard, P.M., Vigre, H., Thamsborg, S.M. and Lind, P. 2003. Cryptosporidium parvum, infectivity and pathogenicity of the “porcine” genotype. Parasitology 126, 407–416. Fayer, R. 1994. Effect of high temperature on infectivity of Cryptosporidium parvum oocysts in water. Appl. Environ. Microbiol. 60, 2732–2735. Fayer, R. 1995. Effect of sodium hypoclorite exposure on infectivity of Cryptosporidium parvum oocysts for neonatal BALB/c mice. Appl. Environ. Microbiol. 61, 844–846. Fayer, R. 1997. The general biology of Cryptosporidium. In Cryptosporidium and Cryptosporidiosis, Fayer, R. Ed., CRC Press, Boca Raton, FL, pp. 1–41. Fayer, R., Dubey, J.P. and Lindsay, D.S. 2004. Zoonotic protozoa, from land to sea. Trends Parasitol. 20, 531–536. Fayer, R., Graczyk, T. K., Cranfield, M. R. and Trout, J. 1996. Effect of low molecular weight gases on disinfection of Cryptosporidium oocysts. Appl. Environ. Microbiol. 62, 3908–3909. Fayer, R. and Nerad, T. 1996. Effects of low temperatures on viability of Cryptosporidium parvum oocysts. Appl. Environ. Microbiol. 62, 1431–1433. Fayer, R., Nerad, T., Rall, W., Lindsay, D.S. and Blagburn, B.L. 1991. Studies on cryopreservation of Cryptosporidium parvum. J. Parasitol. 77, 357–361. Fayer, R., Santin, M. and Xiao, L. 2005. Cryptosporidium bovis n. sp. (Apicomplexa, Cryptosporidiidae) in cattle (Bos taurus). J. Parasitol. 91, 624–629. Fayer, R., Speer, C.A. and Dubey, J.P. 1990. General biology of Cryptosporidium, in Cryptosporidiosis of Man and Animals, Dubey, J.P., Speer, C.A. and Fayer, R., Eds., CRC Press, Boca Raton, FL, pp. 1–29. 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. Fayer, R., Trout, J.M., Walsh, E. and Cole, R. 2000. Rotifers ingest oocysts of Cryptosporidium parvum. J. Eukaryot. Microbiol. 47, 161–163. Fayer, R., Trout, J.M., Xiao, L., Morgan, U., Lal, A.A. and Dubey, J.P. 2001. Cryptosporidium canis n. sp. from domestic dogs. J. Parasitol. 87, 1415–1422. Fleta, J., Sanchez-Acedo, C., Clavel, A. and Quilez, J. 1995. Detection of Cryptosporidium oocysts in extraintestinal tissues of sheep and pigs. Vet. Parasitol. 59, 201–205. Gatei, W., Greensill, J., Ashford, R.W., Cuevas, L.E., Parry, C.M., Cunliffe, N.A., Beeching, N.J. and Hart, C.A. 2003. Molecular analysis of the 18S rRNA gene of Cryptosporidium parasites from patients with or without human immunodeficiency virus infections living in Kenya, Malawi, Brazil, the United Kingdom, and Vietnam. J. Clin. Microbiol. 41, 1458–1462. Gibson, S.V. and Wagner, J.E. 1986. Cryptosporidiosis in guinea pigs, a retrospective study. J. Am. Vet. Med. Assoc. 189, 1033–1034. Girouard, D., Gallant, J., Akiyoshi, D.E., Nunnari, J. and Tzipori, S. 2006. Failure to propagate Cryptosporidium spp. in cell-free culture. J. Parasitol. 92, 399–400. Glaberman, S., Sulaiman, I.M., Bern, C., Limor, J., Peng, M.M., Morgan, U., Gilman, R., Lal, A.A. and Xiao, L. 2001. A multilocus genotypic analysis of Cryptosporidium meleagridis. J. Eukaryot. Microbiol. (Suppl.) 19–22S. Graczyk, T.K., Fayer, R. and Cranfield, M.R. 1996. Cryptosporidium parvum is not transmissible to fish, amphibia, or reptiles. J. Parasitol. 82, 748–751. Grimason, A., Smith, H., Thitai, W., Smith, P., Jackson, M. and Girdwood, R. 1993. Occurrence and removal of Cryptosporidium spp. oocysts and Giardia spp. cysts in Kenyan waste stabilisation ponds. Wat. Sci. Tech. 27, 97–104. Guyot, K., Follet-Dumoulin, A., Recort, C., Lelievre, C., Cailliez, J.C. and Dei-Cas, E. 2001. Molecular characterization of Cryptosporidium isolates obtained from humans in France. J. Clin. Microbiol. 39, 3472–3480.


38


Hanes D.E., Worobo, R.W., Orlandi, P.A., Burr, D.H., Miliotis, M.D., Robi, M.G., Bier, J.W., Arrowood, M.J., Churey, J.J. and Jackson, G.J. 2002. Inactivation of Cryptosporidium parvum oocysts in fresh apple cider by UV irradiation. Appl. Environ. Microbiol. 68, 4168–4172. Harp, J.A., Fayer, R., Pesch, B.A. and Jackson, G.J. 1996. Effect of pasteurization on infectivity of Cryptosporidium parvum oocysts in water and milk. Appl. Environ. Microbiol. 62, 2866–2868. Harris, J.R. and Petry, F. 1999. Cryptosporidium parvum, structural components of the oocyst wall. J. Parasitol. 85, 839–849. Harris, J.R., Adrian, M. and Petry, F. 2003. Structure of the Cryptosporidium microneme, a metabolically and osmotically labile organelle. Micron 34, 65–78. Harris, J.R., Adrian, M. and Petry, F. 2004. Amylopectin, a major constituent of the residual body in Cryptosporidium oocysts. Parasitology 128, 269–282. Hijjawi, N.S., Meloni, B.P., Ng’anzo, M., Ryan, U.M., Olsen, M.E., Cox, P.T., Monis, P.T. and Thompson, R.C. 2004. Complete development of Cryptosporidium parvum in host cell-free culture. Int. J. Parasitol. 34, 769–777. Hijjawi, N.S., Meloni, B.P., Ryan, U.M., Olson, M.E. and Thompson, R.C. 2002. Successful in vitro cultivation of Cryptosporidium andersoni, evidence for the existence of novel extracellular stages in the life cycle and implications for the classification of Cryptosporidium. Int. J. Parasitol. 32, 1719–1726. Holton, J., Nye, P. and McDonald V. 1994. Efficacy of selected disinfectants against mycobacteria and cryptosporidia. J. Hosp. Infect. 27, 105–115. Hoover, D.M., Hoerr, F.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. Huang, B.Q., Chen, X.-M. and LaRusso, N.F. 2004. Cryptosporidium parvum attachment to and internalization by human biliary epithelia in vitro, a morphological study. J. Parasitol. 90, 212–221. Inman, L.R. and Takeuchi, A. 1979. Spontaneous cryptosporidiosis in an adult female rabbit. Vet. Pathol. 16, 89–95. Iseki, M. 1979. Cryptosporidium felis sp. n. (Protozoa, Eimeriorina) from the domestic cat. Jpn. J. Parasitol. 28, 285–307. Jenkins, M.B., Bowman, D.D. and Ghiorse, W.C. 1998. Inactivation of Cryptosporidium parvum oocysts by ammonia. Appl. Environ. Microbiol. 64, 784–788. Jenkins, M.C., Higgins, J., Kniel., K., Trout, M.J. and Fayer, R. 2004. Protection of calves against cryptosporidiosis by oral inoculation with gamma-irradiated Cryptosporidium parvum oocysts. J. Parasitol. 90, 830–833. Jenkins, M.C., Trout, M.J. and Fayer, R. 1998a. Development and application of an improved semiquantitative technique for detecting low-level Cryptosporidium parvum infections in mouse tissue using polymerase chain reaction. J. Parasitol. 84, 182–186. Jenkins, M.C., Trout, M.J., Higgins, J., Dorsch, M., Veal, D. and Fayer, R. 2003. Comparison of tests for viable and infectious Cryptosporidium parvum oocysts. Parasitol. Res. 89, 1–5. Johnson, A.M., Linden, K., Ciociola, K.M., DeLeon, R., Widmer, G. and Rochelle, P. 2005. UV inactivation of Cryptosporidium hominis measured in cell culture. Appl. Environ. Microbiol. 71, 2800–2802. Karim, M.R., Manshadi, F.D., Kariscak, M.M. and Gerba, C.P. 2004. The persistence and removal of enteric pathogens in constructed wetlands. Wat. Res. 38, 1831–1837. Kato, S., Jenkins, M.B., Fogarty, E.A. and Bowman, D.D. 2002. Effects of freeze-thaw events on the viability of Cryptosporidium parvum oocysts in soil. J. Parasitol. 88, 718–722. Keegan, A.R., Fanok, S., Monis, P.T. and Saint, C.P. 2003. Cell culture-Taqman PCR assay for evaluation of Cryptosporidium parvum disinfection. Appl. Environ. Microbiol. 69, 2505–2511. Keithly, J.S., Langreth, S.G., Buttle, K.F. and Mannella, C.A. 2005. Electron tomographic and ultrastructural analysis of the Cryptosporidium parvum relict mitochondrion, its associated membranes, and organelles. J. Eukaryot. Microbiol. 52, 132–140. King B.J., Keegan, A.R., Monis, P.T. and Saint, C.P. 2005. Experimental temperature controls Cryptosporidium oocyst metabolic rate and associated retention of infectivity. Appl. Environ. Microbiol. 71, 3848–3857. Korich, D.G., Mead, J.R., Madore, M.S., Sinclair, N.A. and Sterling, C.R. 1990. Effects of ozone, chlorine dioxide, chlorine, and monochloramine on Cryptosporidium parvum oocyst viability. Appl. Environ. Microbiol. 56, 1423–1428. Koudela, B. and Modry, D. 1998. New species of Cryptosporidium (Apicomplexa, Cryptosporidiidae) from lizards. Fol. Parasitol. 45, 93–100.


General Biology

39

Langer, R.C. and Riggs M.W. 1999. Cryptosporidium parvum apical complex glycoprotein CSL contains a sporozoite ligand for intestinal epithelial cells. Infect. Immun. 67, 5282–5291. Langer, R.C., Schaefer, D.A. and Riggs, M.W. 2001. Characterization of an intestinal epithelial cell receptor recognized by the Cryptosporidium parvum sporozoite ligand CSL. Infect. Immun. 69, 1661–1670. Lasser, K.H., Lewin, K.J. and Ryning, F.W. 1979. Cryptosporidial enteritis in a patient with congenital hypogammaglobulinemia. Hum. Pathol. 10, 234–240. Leander, B.S., Clopton, R.E. and Keeling, P.J. 2003. Phylogeny of gregarines (Apicomplexa) as inferred from small-subunit rDNA and beta-tubulin. Int. J. Syst. Evol. Microbiol. 53, 345–354. Leoni, F., Amar, C., Nichols, G., Pedraza-Diaz, S. and McLauchlin, J. 2006. Genetic analysis of Cryptosporidium from 2414 humans with diarrhoea in England between 1985 and 2000. J. Med. Microbiol. 55, 703–707. Levine, N.D.1961. Protozoan Parasites of Domestic Animals and of Man. Burgess Publishing, Minneapolis, 245. Levine, N.D. 1980. Some corrections of coccidian (Apicomplexa, Protozoa) nomenclature. J. Parasitol. 66, 830–834. Levine, N.D. 1985. Phylum II. Apicomplexa Levine 1970, in Illustrated Guide to the Protozoa, Lee, J.J., Hutner, S.H. and Bovee, E.C., Eds., Society of Protozoologists, Lawrence, Kansas, 322–374. Lindsay, D.S., Upton S.J., Owens, D.S., Morgan, U.M., Mead, J.R. and Blagburn, B.L. 2000. Cryptosporidium andersoni n. sp. (Apicomplexa, Cryptosporiidae) from cattle, Bos taurus. J. Eukaryot. Microbiol. 47, 91–95. Lorenzo-Lorenzo, M.J., Ares-Mazas, M.E., Villa Corta, I. and Duran-Oreiro, D. 1993. Effect of ultraviolet disinfection of drinking water on the viability of Cryptosporidium parvum oocysts. J. Parasitol. 79, 67–70. Lumb, R., Smith, K., O’Donoghue, P.J. and Lanser, J.A. 1988. Ultrastructure of the attachment of Cryptosporidium sporozoites to tissue culture cells. Parasitol. Res. 74, 531–536. MacKenzie, W.R., Hoxie, N.J., Proctor, M.E., Gradus, M.S., Blair, K.A., Peterson, D.E., Kazmierczak, J.J., Addiss, D.G., Fox, K.R., Rose, J.B. and Davis, J.P. 1994. A massive outbreak in Milwaukee of Cryptosporidium infection transmitted through the public water supply. N. Eng. J. Med. 331, 161–167. Matschoulsky, S.N. 1947. Coccidia of wild birds in Buryat. Trudy Buryat-Mongol. Zoovet. Inst. 3, 93–101. [In Russian] Matsui, T. Fujino, T. and Tsuji, M. 1999. Infectivity to hosts of the endogenous stages of chicken and murine Cryptosporidium. J. Vet. Med. Sci. 61, 471–474. McGuigan, K.G., Mendez-Hermida, F., Castro-Hermida, J.K.A., Ares-Mazas, E., Kehoe, S.C., Boyle, M., Sichel, C., Fernandez-Ibanez, P., Meyer, B.P., Ramalingham, S. and Meyer, E.A. 2006. Batch solar disinfection inactivates oocysts of Cryptosporidium parvum and cysts of Giardia muris in drinking water. J. Appl. Microbiol. 101, 453–463. Meisel, J.L., Perera, D.R., Meligro, C. and Rubin, C.E. 1976. Overwhelming watery diarrhea associated with a Cryptosporidium in an immunosuppressed patient. Gastroenterology 70, 1156–1160. Meutin, D.J., Van Kruiningen, H.J. and Kein, D.H. 1974. Cryptosporidiosis in a calf. J. Am. Vet. Med. Assoc. 165, 914–917. Morgan-Ryan, U.M., Fall, A., Ward, L.A., Hijjawi, N., Sulaiman, I., Fayer, R., Thompson, R.C.A, Olsen, M., Lal, A.A. and Xiao, L. 2002. Cryptosporidium hominis n. sp. (Apicomplexa, Cryptosporidiidae) from Homo sapiens. J. Eukaryot. Microbiol. 49, 433–440. Morita, S., Namikoshi, A., Hirata, T., Oguma, K., Katayama, H., Ohgaki, S., Motoyama, N. and Fujiwara, M. 2002. Efficacy of UV irradiation in inactivating Cryptosporidium parvum oocysts. Appl. Environ. Microbiol. 68, 5387–5393. 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. Nokes, R.L., Gerba, C.P. and Karpiscak, M.M. 2003. Microbial water quality improvement by small scale onsite subsurface wetland treatment. J. Environ. Sci. Hlth. A38, 1849–1855. O’Hara, S.P., Huang, B.Q., Chen, X.-M., Nelson, J. and LaRusso, N.F. 2005. Distribution of Cryptosporidium parvum sporozoite apical organelles during attachment to and internalization by cultured biliary epithelial cells. J. Parasitol. 91, 995–999. Ortega, Y.R. and Liao, J. 2006. Microwave inactivation of Cyclospora cayetanensis sporulation and viability of Cryptosporidium parvum oocysts. J. Food Prot. 69, 1957–1960.


40


Oyane, I., Furuta, M., Stavarache, C.E., Hashiba, K., Mukai, S., Nakanishi, M., Kimata, I. and Maeda, Y. 2005. Inactivation of Cryptosporidium parvum by ultrasonic irradiation. Environ. Sci. Technol. 39, 7294–7298. Panciera, R.J., Thomassen, R.W. and Garner, F.M. 1971. Cryptosporidial infection in a calf. Vet. Pathol. 8, 479–484. Paperna, I., Landsberg, J.H. and Ostrovska, K. 1986. Cryptosporidia of lower vertebrates. Soc. Protozool. Abstracts 245. Parker, J. and Smith, H. 1993. Destruction of oocysts of Cryptosporidium parvum by sand and chlorine. Wat. Res. 27, 729–731. Pavlasek, I. 1984. Effect of disinfectants in infectiousness of oocysts of Cryptosporidium sp. Cs. Epidem. Mikrobiol. Immunol. 33, 97–101. Pavlasek, I. 1999. Cryptosporidia, biology diagnosis, host spectrum, specificity and the environment. Remedia Klin. Microbiol. 3, 290–302. [In Czech] 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. Pedraza-Diaz, S., Amar, C. and McLauchlin, J. 2000. The identification and characterisation of an unusual genotype of Cryptosporidium from human faeces as Cryptosporidium meleagridis. FEMS Microbiol Lett. 189, 189–194. Peeters, J. E., Ares Mazas, E., Masschelein, W. J., Villacorta, I. and Debacker, E. 1989. Effect of disinfection of drinking water with ozone or chlorine dioxide on survival of Cryptosporidium parvum oocysts. Appl. Environ. Microbiol. 55, 1519–1522. 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. Putignani, L. 2005. The unusual architecture and predicted function of the mitochondrion organelle in Cryptosporidium parvum and hominis species, the strong paradigm of the structure-function relationship. Parasitology 47, 217–225. Qadripur, S.A. and Klose, L. 1985. Cryptosporidium enteritidis. Occurrence in southern Lower Saxony: Study of urticaria as an example. Dermatol. Monatsschr. 171, 438–442. Quilez, J., Sanchez-Acedo, C., Avendano, C., delCacho, E. and Lopez-Berna, F. 2005. Efficacy of two peroxygen-based disinfectants for inactivation of Cryptosporidium parvum oocysts, Appl. Environ. Microbiol. 71, 2479–2483. Quinn, C.M. and Betts, W.B. 1993. Longer term viability status of chlorine-treated Cryptosporidium oocysts in tap water. Biomed. Lett. 48, 315–318. Ransome, M.E., Whitmore, T.N. Carrington, E.G. 1993. Effect of disinfectants on the viability of Cryptosporidium parvum oocysts. Wat. Supply 11, 103–117. Reduker, D.W., Speer, C.A. and Blixt, J.A. 1985. Ultrastructure of Cryptosporidium parvum oocysts and excysting sporozoites as revealed by high pressure scanning electron microscopy. J. Protozool. 32, 708–711. Riggs, M.W., Stone, A.L., Yount, P.A., Langer, R.C., Arrowood, M.J. and Bentley, D. L. 1997. Protective monoclonal antibody defines a circumsporozoite-like glycoprotein exoantigen of Cryptosporidium parvum sporozoites and merozoites. J. Immunol. 158, 1787–1795. Robertson, L.J., Campbell, A.T. and Smith, H.V. 1992. Survival of Cryptosporidium parvum oocysts under various environmental pressures. Appl. Environ. Microbiol. 58, 3494–3500. Rosales, M.J., Cordon, G.P., Moreno, M.S. and Sanchez, C.M. 2005. Extracellular llike-gregarine stages of Cryptosporidium parvum. Acta Trop. 95, 74–78. Ryan, U.M., Bath, C., Robertson, I., Read, C., Elliot, A., Mcinnes, L., Traub, R. and Besier, B. 2005. Sheep may not be an important zoonotic reservoir for Cryptosporidium and Giardia parasites. Appl. Environ. Microbiol. 71, 4992–4997. 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. Ryan, U.M., Xiao, L., Read, C., Sulaiman, I.M., Monis, P., Lal, A.A., Fayer, R., and Pavlasek, I. 2003. A redescription of Cryptosporidium galli Pavlasek, 1999 (Apicomplexa, Cryptosporidiidae) from birds. J. Parasitol. 89, 809–813.


General Biology

41

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.


42


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.


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.

43


44

II. A.


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


Genomics

45

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


46


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,


Genomics

47

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


48

IV.


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.


Genomics

49

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


50


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


Genomics

51

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


52


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


Genomics

53

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.

References Abrahamsen, M.S., Schroeder, A.A., and Lancto, C.A. 1996. Differential mRNA display analysis of gene expression in Cryptosporidium parvum infected HCT-8 cells. J. Eukaryot. Microbiol. 43, 80 and 81s. Abrahamsen, M.S., Templeton, T.J., Enomoto, S., Abrahante, J.E., Zhu, G., Lancto, C.A., Deng, M., Liu, C., Widmer, G., Tzipori, S., Buck, G.A., Xu, P., Bankier, A.T., Dear, P.H., Konfortov, B.A., Spriggs, H.F., Iyer, L., Anantharaman, V., Aravind, L., and Kapur, V. 2004. Complete genome sequence of the apicomplexan, Cryptosporidium parvum. Science 304, 441–445. Ajioka, J.W., Boothroyd, J.C., Brunk, B.P., Hehl, A., Hillier, L., Manger, I.D., Marra, M., Overton, G.C., Roos, D.S., Wan, K.L., Waterston, R., and Sibley, L.D. 1998. Gene discovery by EST sequencing in Toxoplasma gondii reveals sequences restricted to the Apicomplexa. Genome Res. 8, 18–28. Aurrecoechea, C.M., Heiges, H., Wang, Z., Wang, S., Fischer, P., Rhodes, J., Miller, Kraemer, E., Stoeckert, C.J., Roos, D.S., and Kissinger, J.C. 2007. ApiDB: Integrated Resources for the Apicomplexan Bioinformatics Resource Center. Nucleic Acids Res. 35: D427–D430. Bankier, A.T., Spriggs, H.F., Fartmann, B., Konfortov, B.A., Madera, M., Vogel, C., Teichmann, S.A., Ivens, A.and Dear, P.H., 2003. Integrated mapping, chromosomal sequencing and sequence analysis of Cryptosporidium parvum. Genome Res. 13, 1787–1799. Barta, J.R. and Thompson, R.C. 2006. What is Cryptosporidium? Reappraising its biology and phylogenetic affinities. Trends Parasitol. 22, 463–468. Blunt, D.S., Khramtsov, N.V., Upton, S.J., and Montelone, B.A. 1997. Molecular karyotype analysis of Cryptosporidium parvum: evidence for eight chromosomes and a low-molecular-size molecule. Clin. Diagn. Lab. Immunol. 4, 11–13. Caccio, S., Camilli, R., La Rosa, G., and Pozio., E. 1998. Establishing the Cryptosporidium parvum karyotype by NotI and SfiI restriction analysis and Southern hybridization. Gene 219, 73–79. Cama, V.A., Arrowood, M.J., Ortega, Y.R., and Xiao, L.. 2006. Molecular characterization of the Cryptosporidium parvum IOWA isolate kept in different laboratories. J. Eukaryot. Microbiol. 53, S40–42. Carlton, J.M., Angiuoli, S.V., Suh, B.B., Kooij, T.W., Pertea, M., Silva, J.C., Ermolaeva, M.D., Allen, J.E., Selengut, J.D., Koo, H.L., Peterson, J.D., Pop, M., Kosack, D.S., Shumway, M.F., Bidwell, S.L., Shallom, S.J., van Aken, S.E., Riedmuller, S.B., Feldblyum, T.V., Cho, J.K., Quackenbush, J., Sedegah, M., Shoaibi, A., Cummings, L.M., Florens, L., Yates, J.R., Raine, J.D., Sinden, R.E., Harris, M.A., Cunningham, D.A., Preiser, P.R., Bergman, L.W., Vaidya, A.B., van Lin, L.H., Janse, C.J., Waters, A.P., Smith, H.O., White, O.R., Salzberg, S.L., Venter, J.C., Fraser, C.M., Hoffman, S.L., Gardner, M.J., and Carucci, D.J. 2002. Genome sequence and comparative analysis of the model rodent malaria parasite Plasmodium yoelii yoelii. Nature 419, 512–519. Carreno, R.A., Martin, D.S., and Barta, J.R. 1999. Cryptosporidium is more closely related to the gregarines than to coccidia as shown by phylogenetic analysis of apicomplexan parasites inferred using small-subunit ribosomal RNA gene sequences. Parasitol. Res. 85, 899–904. Deng, M., Templeton, T.J., London, N.R., Bauer, C., Schroeder, A.A., and Abrahamsen, M.S. 2002. Cryptosporidium parvum genes containing thrombospondin type 1 domains. Infect. Immun. 70, 6987–6895.


54


Dzierszinski, F., Popescu, O., Toursel, C., Slomianny, C., Yahiaoui, B., and Tomavo, S. 1999. The protozoan parasite Toxoplasma gondii expresses two functional plant-like glycolytic enzymes: Implications for evolutionary origin of apicomplexans. J. Biol. Chem. 274, 24888–24895. Fayer, R. 1997. Cryptosporidium and Cryptosporidiosis, CRC Press, Boca Raton, FL, 270 pp. Foth, B.J., Ralph, S.A., Tonkin, C.J., Struck, N.S., Fraunholz, M., Roos, D.S., Cowman, A.F. and McFadden, G.I. 2003. Dissecting apicoplast targeting in the malaria parasite Plasmodium falciparum. Science 299, 705–708. Gardner, M.J., Bishop, R., Shah, T., de Villiers, E.P., Carlton, J.M., Hall, N., Ren, Q., Paulsen, I.T., Pain, A., Berriman, M., Wilson, R.J., Sato, S., Ralph, S.A., Mann, D.J., Xiong, Z., Shallom, S.J., Weidman, J., Jiang, L., Lynn, J., Weaver, B., Shoaibi, A., Domingo, A.R., Wasawo, D., Crabtree, J., Wortman, J.R., Haas, B., Angiuoli, S.V., Creasy, T.H., Lu, C., Suh, B., Silva, J.C., Utterback, T.R., Feldblyum, T.V., Pertea, M., Allen, J., Nierman, W.C., Taracha, E.L., Salzberg, S.L., White, O.R., Fitzhugh, H.A., Morzaria, S., Venter, J.C., Fraser, C.M., and Nene, V. 2005. Genome sequence of Theileria parva, a bovine pathogen that transforms lymphocytes. Science 309, 134–137. Gardner, M.J., Hall, N., Fung, E., White, O., Berriman, M., Hyman, R.W., Carlton, J.M., Pain, A., Nelson, K.E., Bowman, S., Paulsen, I.T., James, K., Eisen, J.A., Rutherford, K., Salzberg, S.L., Craig, A., Kyes, S., Chan, M.S., Nene, V., Shallom, S.J., Suh, B., Peterson, J., Angiuoli, S., Pertea, M., Allen, J., Selengut, J., Haft, D., Mather, M.W., Vaidya, A.B., Martin, D.M., Fairlamb, A.H., Fraunholz, M.J., Roos, D.S., Ralph, S.A., McFadden, G.I., Cummings, L.M., Subramanian, G.M., Mungall, C., Venter, J.C., Carucci, D.J., Hoffman, S.L., Newbold, C., Davis, R.W., Fraser, C.M., and Barrell, B. 2002. Genome sequence of the human malaria parasite Plasmodium falciparum. Nature 419, 498–511. Hays, M.P., Mosier, D.A., and Oberst, R.D. 1995. Enhanced karyotype resolution of Cryptosporidium parvum by contour-clamped homogeneous electric fields. Vet. Parasitol. 58, 273–280. Howe, D.K. 2001. Initiation of a Sarcocystis neurona expressed sequence tag (EST) sequencing project: A preliminary report. Vet. Parasitol. 95, 233–239. Huang, J., Mullapudi, N., Lancto, C.A., Scott, M., Abrahamsen, M.S., and Kissinger, J.C.. 2004a. Phylogenomic evidence supports past endosymbiosis, intracellular and horizontal gene transfer in Cryptosporidium parvum. Genome Biol. 5, R88. Huang, J., Mullapudi, N., Sicheritz-Ponten, T., and Kissinger, J.C. 2004b. A first glimpse into the pattern and scale of gene transfer in Apicomplexa. Int. J. Parasitol. 34, 265–274. 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, New York. Levine, N.D. 1970. Taxonomy of the Sporozoa. J. Protozool. 56, 208–209. Levine, N.D. 1988. The Protozoan Phylum Apicomplexa Vol. I. CRC Press, Boca Raton, FL. Li, L., Brunk, B.P., Kissinger, J.C., Pape, D., Tang, K., Cole, R.H., Martin, J., Wylie, T., Dante, M., Fogarty, S.J., Howe, D.K., Liberator, P., Diaz, C., Anderson, J., White, M., Jerome, M.E., Johnson, E.A., Radke, J.A., Stoeckert, Jr. C.J., Waterston, R.H., Clifton, S.W., Roos, D.S., and Sibley. L.D. 2003a. Gene discovery in the apicomplexa as revealed by EST sequencing and assembly of a comparative gene database. Genome Res. 13, 443–454. Li, L., Stoeckert, C.J., and Roos, D.S. 2003b. OrthoMCL: Identification of ortholog groups for eukaryotic genomes. Genome Res. 13, 2178–2189. Liu, C., Vigdorovich, V., Kapur, V., and Abrahamsen, M.S. 1999. A random survey of the Cryptosporidium parvum genome. Infect. Immun. 67, 3960–3969. Madern, D., Cai, X., Abrahamsen, M.S., and Zhu, G. 2004. Evolution of Cryptosporidium parvum lactate dehydrogenase from malate dehydrogenase by a very recent event of gene duplication. Mol. Biol. Evol. 21, 489–497. Mead, J.R., Arrowood, M.J., Current, W.L., and Sterling, C.R. 1988. Field inversion gel electrophoretic separation of Cryptosporidium spp. chromosome-sized DNA. J. Parasitol. 74, 366–369. Morgan, U.M., Xiao, L., Fayer, R., Graczyk, T.K., Lal, A.A., Deplazes, P., and Thompson, R.C. 1999a. Phylogenetic analysis of Cryptosporidium isolates from captive reptiles using 18S rDNA sequence data and random amplified polymorphic DNA analysis. J. Parasitol. 85, 525–550. Morgan, U.M., Xiao, L., Fayer, R., Lal, A.A., and Thompson, R.C. 1999b. Variation in Cryptosporidium: Towards a taxonomic revision of the genus. Int. J. Parasitol. 29, 1733–1751.


Genomics

55

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.


56


Umejiego, N.N., Li, C., Riera, T., Hedstrom, L., and Striepen, B. 2004. Cryptosporidium parvum IMP dehydrogenase: identification of functional, structural, and dynamic properties that can be exploited for drug design. J. Biol. Chem. 279, 40320–40327. Vedadi, M., Lew, J., Artz, J., Amani, M., Zhao, Y., Dong, A., Wasney, G.A., Gao, M., Hills, T., Brokx, S., Qiu, W., Sharma, S., Diassiti, A., Alam, Z., Melone, M., Mulichak, A., Wernimont, A., Bray, J., Loppnau, P., Plotnikova, O., Newberry, K., Sundararajan, E., Houston, S., Walker, J., Tempel, W., Bochkarev, A., Kozieradzki, I., Edwards, A., Arrowsmith, C., Roos, D., Kain, K., and Hui. R. 2007. Genome-scale protein expression and structural biology of Plasmodium falciparum and related apicomplexan organisms. Mol. Biochem. Parasitol. 151, 100–110. Wan, K. L., Chong, S.P., Ng, S.T., Shirley, M.W., Tomley, F.M., and Jangi. M.S. 1999. A survey of genes in Eimeria tenella merozoites by EST sequencing. Int. J. Parasitol. 29, 1885–1892. Widmer, G., Lin, L., Kapur, V., Feng, X., and Abrahamsen, M.S. 2002. Genomics and genetics of Cryptosporidium parvum: The key to understanding cryptosporidiosis. Microbes Infect. 4, 1081–1090. Xiao, L., Escalante, L., Yang, C., Sulaiman, I., Escalante, A.A., Montali, R.J., Fayer, R. and Lal, A.A. 1999a. Phylogenetic analysis of Cryptosporidium parasites based on the small-subunit rRNA gene locus. Appl. Environ. Microbiol. 65, 1578–1583. 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., Morgan, U.M., Limor, J., Escalante, A., Arrowood, M., Shulaw, W., Thompson, R.C., Fayer, R., and Lal, A.A. 1999b. Genetic diversity within Cryptosporidium parvum and related Cryptosporidium species. Appl. Environ. Microbiol. 65, 3386–3391. 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. 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., Marchewka, M.J., and Keithly, J.S. 2000b. Cryptosporidium parvum appears to lack a plastid genome. Microbiol. Mol. Biol. Rev. 146, 315–321.


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.

57


58


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


Biochemistry

59

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;


60

FIGURE 3.1



Biochemistry

61

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.


62

IV.


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


Biochemistry

63

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


64


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


Biochemistry

65

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


66


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


Biochemistry

67

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


68


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


Biochemistry

69

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


70


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


Biochemistry

71

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).

References Abelian, A., Walsh, A.P., Lentzen, G., Aboul-Ela, F., and Gait, M.J. 2004. Targeting the A site RNA of the Escherichia coli ribosomal 30 S subunit by 2′-O-methyl oligoribonucleotides: A quantitative equilibrium dialysis binding assay and differential effects of aminoglycoside antibiotics. Biochem. J. 383, 201–208. Abrahamsen, M.S. 1999. Cryptosporidium parvum gene discovery. Adv. Exp. Med. Biol. 473, 241–247. Abrahamsen, M.S., Templeton, T.J., Enomoto, S., Abrahante, J.E., Zhu, G., Lancto, C.A., Deng, M., Liu, C., Widmer, G., Tzipori, S., Buck, G.A., Xu, P., Bankier, A.T., Dear, P.H., Konfortov, B.A., Spriggs, H.F., Iyer, L., Anantharaman, V., Aravind, L., and Kapur, V. 2004. Complete genome sequence of the apicomplexan, Cryptosporidium parvum. Science 304, 441–445. 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. Al-Adhami, B.H., Nichols, R.A., Kusel, J.R., O’Grady, J., and Smith, H.V. 2007. Detection of UV-induced thymine dimers in individual Cryptosporidium parvum and Cryptosporidium hominis oocysts by immunofluorescence microscopy. Appl. Environ. Microbiol. 37, 307–316. Anderson, A.C. 2005a. Two crystal structures of dihydrofolate reductase-thymidylate synthase from Cryptosporidium hominis reveal protein-ligand interactions including a structural basis for observed antifolate resistance. Acta Crystallograph. Sect F Struct. Biol. Cryst. Commun. 61, 258–262. Anderson, A.C. 2005b. Targeting DHFR in parasitic protozoa. Drug Discov. Today 10, 121–128. Bacchi, C.J. and Yarlett, N. 2002. Polyamine metabolism as chemotherapeutic target in protozoan parasites. Mini Rev. Med. Chem. 2, 553–563. Baum, J., Richard, D., Healer, J., Rug, M., Krnajski, Z., Gilberger, T.W., Green, J.L., Holder, A.A., and Cowman, A.F. 2006. A conserved molecular motor drives cell invasion and gliding motility across malaria life cycle stages and other apicomplexan parasites. J. Biol. Chem. 281, 5197–5208. Birkholtz, L.M., Wrenger, C., Joubert, F., Wells, G.A., Walter, R.D., and Louw, A.I. 2004. Parasite-specific inserts in the bifunctional S-adenosylmethionine decarboxylase/ornithine decarboxylase of Plasmodium falciparum modulate catalytic activities and domain interactions. Biochem. J. 377, 439–448. Bonnin, A., Dubremetz, J.F., and Camerlynck, P. 1991. Characterization and immunolocalization of an oocyst wall antigen of Cryptosporidium parvum (Protozoa: Apicomplexa). Parasitology 103, 171–177. Bonnin, A., Ojcius, D.M., Souque, P., Barnes, D.A., Doyle, P.S., Gut, J., Nelson, R.G., Petersen, C., and Dubremetz, J.F. 2001. Characterization of a monoclonal antibody reacting with antigen-4 domain of gp900 in Cryptosporidium parvum invasive stages. Parasitol. Res. 87, 589–592.


72


Brophy, V.H., Vasquez, J., Nelson, R.G., Forney, J.R., Rosowsky, A., and Sibley, C.H. 2000. Identification of Cryptosporidium parvum dihydrofolate reductase inhibitors by complementation in Saccharomyces cerevisiae. Antimicrob. Agents Chemother. 44, 1019–1028. Brown, G.W., Hines, J.C., Fisher, P., and Ray, D.S. 1994. Isolation of the genes encoding the 51-kilodalton and 28-kilodalton subunits of Crithidia fasciculata replication protein A. Mol. Biochem. Parasitol. 63, 135–142. Caccio, S., La Rosa, G., and Pozio, E. 1997. The beta-tubulin gene of Cryptosporidium parvum. Mol. Biochem. Parasitol. 89, 307–311. Cai, X., Lancto, C.A., Abrahamsen, M.S., and Zhu, G. 2004. Intron-containing beta-tubulin transcripts in Cryptosporidium parvum cultured in vitro. Microbiology 150, 1191–1195. Cai, X., Herschap, D., and Zhu, G. 2005. Functional characterization of an evolutionarily distinct phosphopantetheinyl transferase in the apicomplexan Cryptosporidium parvum. Eukaryot. Cell 4, 1211–1220. Camero, L., Arrowood, M., Shulaw, W., Lal, A.A., and Xiao, L. 1999. Characterization of new monoclonal antibodies against Cryptosporidium parvum sporozoites. J. Eukaryot. Microbiol. 46, 58S–59S. Carreno, R.A., Martin, D.S., and Barta, J.R. 1999. Cryptosporidium is more closely related to the gregarines than to coccidia as shown by phylogenetic analysis of apicomplexan parasites inferred using small-subunit ribosomal RNA gene sequences. Parasitol. Res. 85, 899–904. Cevallos, A.M., Zhang, X., Waldor, M.K., Jaison, S., Zhou, X., Tzipori, S., Neutra, M.R., and Ward, H.D. 2000. Molecular cloning and expression of a gene encoding Cryptosporidium parvum glycoproteins gp40 and gp15. Infect. Immun. 68, 4108–4116. Chen, X.M., O’Hara, S.P., Huang, B.Q., Nelson, J.B., Lin, J.J., Zhu, G., Ward, H.D., and LaRusso, N.F. 2004. Apical organelle discharge by Cryptosporidium parvum is temperature, cytoskeleton, and intracellular calcium dependent and required for host cell invasion. Infect. Immun. 72, 6806–6816. Coombs, G.H. and Muller, S. 2002. Recent advances in the search for new anti-coccidial drugs. Int. J. Parasitol. 32, 497–508. Ctrnacta, V., Ault, J.G., Stejskal, F., and Keithly, J.S. 2006. Localization of pyruvate: NADP+ oxidoreductase in sporozoites of Cryptosporidium parvum. J. Eukaryot. Microbiol. 53, 225–231. Deng, M., Templeton, T.J., London, N.R., Bauer, C., Schroeder, A.A., and Abrahamsen, M.S. 2002. Cryptosporidium parvum genes containing thrombospondin type 1 domains. Infect. Immun. 70, 6987–6995. Denton, H., Brown, S.M., Roberts, C.W., Alexander, J., McDonald, V., Thong, K.W., and Coombs, G.H. 1996. Comparison of the phosphofructokinase and pyruvate kinase activities of Cryptosporidium parvum, Eimeria tenella and Toxoplasma gondii. Mol. Biochem. Parasitol. 76, 23–29. Dolo, A., Modiano, D., Doumbo, O., Bosman, A., Sidibe, T., Keita, M.M., Naitza, S., Robson, K.J., and Crisanti, A. 1999. Thrombospondin related adhesive protein (TRAP), a potential malaria vaccine candidate. Parassitologia 41, 425–428. Entrala, E. and Mascaro, C. 1997. Glycolytic enzyme activities in Cryptosporidium parvum oocysts. FEMS Microbiol. Lett. 151, 51–57. Fritzler, J.M. and Zhu, G. 2007. Functional characterization of the acyl-[acyl carrier protein] ligase in the Cryptosporidium parvum giant polyketide synthase. Int. J. Parasitol. (In press). Galazka, J., Striepen, B., and Ullman, B. 2006. Adenosine kinase from Cryptosporidium parvum. Mol. Biochem. Parasitol. 149, 223–230. Gardner, M.J., Hall, N., Fung, E., White, O., Berriman, M., Hyman, R.W., Carlton, J.M., Pain, A., Nelson, K.E., Bowman, S., Paulsen, I.T., James, K., Eisen, J.A., Rutherford, K., Salzberg, S.L., Craig, A., Kyes, S., Chan, M.S., Nene, V., Shallom, S.J., Suh, B., Peterson, J., Angiuoli, S., Pertea, M., Allen, J., Selengut, J., Haft, D., Mather, M.W., Vaidya, A.B., Martin, D.M., Fairlamb, A.H., Fraunholz, M.J., Roos, D.S., Ralph, S.A., McFadden, G.I., Cummings, L.M., Subramanian, G.M., Mungall, C., Venter, J.C., Carucci, D.J., Hoffman, S.L., Newbold, C., Davis, R.W., Fraser, C.M., and Barrell, B. 2002. Genome sequence of the human malaria parasite Plasmodium falciparum. Nature 419, 498–511. Gooze, L., Kim, K., Petersen, C., Gut, J., and Nelson, R.G. 1991. Amplification of a Cryptosporidium parvum gene fragment encoding thymidylate synthase. J. Protozool. 38, 56S–58S. Gordon, J.L. and Sibley, L.D. 2005. Comparative genome analysis reveals a conserved family of actin-like proteins in apicomplexan parasites. BMC Genomics 6, 179. Gornicki, P. 2003. Apicoplast fatty acid biosynthesis as a target for medical intervention in apicomplexan parasites. Int. J. Parasitol. 33, 885–896.


Biochemistry

73

Heiges, M., Wang, H., Robinson, E., Aurrecoechea, C., Gao, X., Kaluskar, N., Rhodes, P., Wang, S., He, C.Z., Su, Y., Miller, J., Kraemer, E., and Kissinger, J.C. 2006. CryptoDB: A Cryptosporidium bioinformatics resource update. Nucleic Acids Res. 34, D419–422. Huang, J., Mullapudi, N., Lancto, C.A., Scott, M., Abrahamsen, M.S., and Kissinger, J.C. 2004. Phylogenomic evidence supports past endosymbiosis, intracellular and horizontal gene transfer in Cryptosporidium parvum. Genome Biol. 5, R88. Iochmann, S., Sagodira, S., Mevelec, M.N., Reperant, J.M., Naciri, M., Coursaget, P., and Bout, D. 1999. Comparison of the humoral and cellular immune responses to two preparations of Cryptosporidium parvum CP15/60 recombinant protein. Microb. Pathog. 26, 307–315. Jelenska, J., Crawford, M.J., Harb, O.S., Zuther, E., Haselkorn, R., Roos, D.S., and Gornicki, P. 2001. Subcellular localization of acetyl-CoA carboxylase in the apicomplexan parasite Toxoplasma gondii. Proc. Natl. Acad. Sci. U. S. A. 98, 2723–2728. Jelenska, J., Sirikhachornkit, A., Haselkorn, R., and Gornicki, P. 2002. The carboxyltransferase activity of the apicoplast acetyl-CoA carboxylase of Toxoplasma gondii is the target of aryloxyphenoxypropionate inhibitors. J. Biol. Chem. 277, 23208–23215. Jenkins, M.C., Trout, J., Murphy, C., Harp, J.A., Higgins, J., Wergin, W., and Fayer, R. 1999. Cloning and expression of a DNA sequence encoding a 41-kilodalton Cryptosporidium parvum oocyst wall protein. Clin. Diagn. Lab. Immunol. 6, 912–920. Kabe, Y., Goto, M., Shima, D., Imai, T., Wada, T., Morohashi, K., Shirakawa, M., Hirose, S., and Handa, H. 1999. The role of human MBF1 as a transcriptional coactivator. J. Biol. Chem. 274, 34196–34202. Kappe, S.H., Kaiser, K., and Matuschewski, K. 2003. The Plasmodium sporozoite journey: A rite of passage. Trends Parasitol. 19, 135–143. Kappe, S.H., Buscaglia, C.A., and Nussenzweig, V. 2004. Plasmodium sporozoite molecular cell biology. Annu. Rev. Cell Dev. Biol. 20, 29–59. Kato, S., Lindergard, G., and Mohammed, H.O. 2003. Utility of the Cryptosporidium oocyst wall protein (COWP) gene in a nested PCR approach for detection infection in cattle. Vet. Parasitol. 111, 153–159. Kaul, M., Barbieri, C.M., and Pilch, D.S. 2006. Aminoglycoside-induced reduction in nucleotide mobility at the ribosomal RNA A-site as a potentially key determinant of antibacterial activity. J. Am. Chem. Soc. 128, 1261–1271. Keithly, J.S., Zhu, G., Upton, S.J., Woods, K.M., Martinez, M.P., and Yarlett, N. 1997. Polyamine biosynthesis in Cryptosporidium parvum and its implications for chemotherapy. Mol. Biochem. Parasitol. 88, 35–42. Keithly, J.S., Langreth, S.G., Buttle, K.F., and Mannella, C.A. 2005. Electron tomographic and ultrastructural analysis of the Cryptosporidium parvum relict mitochondrion, its associated membranes, and organelles. J. Eukaryot. Microbiol. 52, 132–140. Khramtsov, N.V., Blunt, D.S., Montelone, B.A., and Upton, S.J. 1996. The putative acetyl-CoA synthetase gene of Cryptosporidium parvum and a new conserved protein motif in acetyl-CoA synthetases. J. Parasitol. 82, 423–427. Khramtsov, N.V., Woods, K.M., Nesterenko, M.V., Dykstra, C.C., and Upton, S.J. 1997. Virus-like, doublestranded RNAs in the parasitic protozoan Cryptosporidium parvum. Mol. Microbiol. 26, 289–300. Khramtsov, N.V. and Upton, S.J. 2000. Association of RNA polymerase complexes of the parasitic protozoan Cryptosporidium parvum with virus-like particles: Heterogeneous system. J. Virol. 74, 5788–5795. Kim, K. and Weiss, L.M. 2004. Toxoplasma gondii: The model apicomplexan. Int. J. Parasitol. 34, 423–432. Kralova, I., Rigden, D.J., Opperdoes, F.R., and Michels, P.A. 2000. Glycerol kinase of Trypanosoma brucei: Cloning, molecular characterization and mutagenesis. Eur. J. Biochem. 267, 2323–2333. Krause, T., Luersen, K., Wrenger, C., Gilberger, T.W., Muller, S., and Walter, R.D. 2000. The ornithine decarboxylase domain of the bifunctional ornithine decarboxylase/S-adenosylmethionine decarboxylase of Plasmodium falciparum: Recombinant expression and catalytic properties of two different constructs. Biochem. J. 352, 287–292. LaGier, M.J., Zhu, G., and Keithly, J.S. 2001. Characterization of a heavy metal ATPase from the apicomplexan Cryptosporidium parvum. Gene 266, 25–34. LaGier, M.J., Keithly, J.S., and Zhu, G. 2002. Characterisation of a novel transporter from Cryptosporidium parvum. Int. J. Parasitol. 32, 877–887. LaGier, M.J., Tachezy, J., Stejskal, F., Kutisova, K., and Keithly, J.S. 2003. Mitochondrial-type iron-sulfur cluster biosynthesis genes (IscS and IscU) in the apicomplexan Cryptosporidium parvum. Microbiology 149, 3519–3530.


74


Lally, N.C., Baird, G.D., McQuay, S.J., Wright, F., and Oliver, J.J. 1992. A 2359-base pair DNA fragment from Cryptosporidium parvum encoding a repetitive oocyst protein. Mol. Biochem. Parasitol. 56, 69–78. Lee, S.H., Stephens, J.L., Paul, K.S., and Englund, P.T. 2006. Fatty acid synthesis by elongases in trypanosomes. Cell 126, 691–699. 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., Gallimore, C.I., Green, J., and McLauchlin, J. 2006. Characterisation of small double stranded RNA molecule in Cryptosporidium hominis, Cryptosporidium felis and Cryptosporidium meleagridis. Parasitol. Int. 55, 299–306. Lilien, R.H., Bailey-Kellogg, C., Anderson, A.C., and Donald, B.R. 2004. A subgroup algorithm to identify cross-rotation peaks consistent with non-crystallographic symmetry. Acta Crystallogr. D Biol. Crystallogr. 60, 1057–1067. Liu, C., Vigdorovich, V., Kapur, V., and Abrahamsen, M.S. 1999. A random survey of the Cryptosporidium parvum genome. Infect. Immun. 67, 3960–3969. Lynch, S.R. and Puglisi, J.D. 2001. Structural origins of aminoglycoside specificity for prokaryotic ribosomes. J. Mol. Biol. 306, 1037–1058. Madern, D., Cai, X., Abrahamsen, M.S., and Zhu, G. 2004. Evolution of Cryptosporidium parvum lactate dehydrogenase from malate dehydrogenase by a very recent event of gene duplication. Mol. Biol. Evol. 21, 489–497. Matsuzaka, T., Shimano, H., Yahagi, N., Yoshikawa, T., Amemiya-Kudo, M., Hasty, A.H., Okazaki, H., Tamura, Y., Iizuka, Y., Ohashi, K., Osuga, J., Takahashi, A., Yato, S., Sone, H., Ishibashi, S., and Yamada, N. 2002. Cloning and characterization of a mammalian fatty acyl-CoA elongase as a lipogenic enzyme regulated by SREBPs. J. Lipid Res. 43, 911–920. McDonald, V., Deer, R.M., Nina, J.M., Wright, S., Chiodini, P.L., and McAdam, K.P. 1991. Characteristics and specificity of hybridoma antibodies against oocyst antigens of Cryptosporidium parvum from man. Parasite Immunol. 13, 251–259. Mead, J.R., Lloyd, R.M., Jr., You, X., Tucker-Burden, C., Arrowood, M.J., and Schinazi, R.F. 1994. Isolation and partial characterization of Cryptosporidium sporozoite and oocyst wall recombinant proteins. J. Eukaryot. Microbiol. 41, 51S. Michalski, W.P., Edgar, J.A., and Prowse, S.J. 1992. Mannitol metabolism in Eimeria tenella. Int. J. Parasitol. 22, 1157–1163. Millership, J.J. and Zhu, G. 2002. Heterogeneous expression and functional analysis of two distinct replication protein A large subunits from Cryptosporidium parvum. Int. J. Parasitol. 32, 1477–1485. Millership, J.J., Cai, X., and Zhu, G. 2004a. Functional characterization of replication protein A2 (RPA2) from Cryptosporidium parvum. Microbiology 150, 1197–1205. Millership, J.J., Waghela, P., Cai, X., Cockerham, A., and Zhu, G. 2004b. Differential expression and interaction of transcription co-activator MBF1 with TATA-binding protein (TBP) in the apicomplexan Cryptosporidium parvum. Microbiology 150, 1207–1213. Mitschler, R.R., Welti, R., and Upton, S.J. 1994. A comparative study of lipid compositions of Cryptosporidium parvum (Apicomplexa) and Madin-Darby bovine kidney cells. J. Eukaryot. Microbiol. 41, 8–12. Moon, Y.A., Shah, N.A., Mohapatra, S., Warrington, J.A., and Horton, J.D. 2001. Identification of a mammalian long chain fatty acyl elongase regulated by sterol regulatory element-binding proteins. J. Biol. Chem. 276, 45358–45366. Morrissette, N.S. and Sibley, L.D. 2002. Cytoskeleton of apicomplexan parasites. Microbiol. Mol. Biol. Rev. 66, 21–38. Muller, S., Da’dara, A., Luersen, K., Wrenger, C., Das Gupta, R., Madhubala, R., and Walter, R.D. 2000. In the human malaria parasite Plasmodium falciparum, polyamines are synthesized by a bifunctional ornithine decarboxylase, S-adenosylmethionine decarboxylase. J. Biol. Chem. 275, 8097–8102. Nina, J.M., McDonald, V., Dyson, D.A., Catchpole, J., Uni, S., Iseki, M., Chiodini, P.L., and McAdam, K.P. 1992. Analysis of oocyst wall and sporozoite antigens from three Cryptosporidium species. Infect. Immun. 60, 1509–1513. O’Connor, R.M., Thorpe, C.M., Cevallos, A.M., and Ward, H.D. 2002. Expression of the highly polymorphic Cryptosporidium parvum Cpgp40/15 gene in genotype I and II isolates. Mol. Biochem. Parasitol. 119, 203–215.


Biochemistry

75

O’Connor, R.M., Kim, K., Khan, F., and Ward, H.D. 2003. Expression of Cpgp40/15 in Toxoplasma gondii: A surrogate system for the study of Cryptosporidium glycoprotein antigens. Infect. Immun. 71, 6027–6034. O’Neil, R.H., Lilien, R.H., Donald, B.R., Stroud, R.M., and Anderson, A.C. 2003. Phylogenetic classification of protozoa based on the structure of the linker domain in the bifunctional enzyme, dihydrofolate reductase-thymidylate synthase. J. Biol. Chem. 278, 52980–52987. Perkins, M.E., Riojas, Y.A., Wu, T.W., and Le Blancq, S.M. 1999. CpABC, a Cryptosporidium parvum ATPbinding cassette protein at the host-parasite boundary in intracellular stages. Proc. Natl. Acad. Sci. U.S.A. 96, 5734–5739. Petersen, C., Gut, J., Doyle, P.S., Crabb, J.H., Nelson, R.G., and Leech, J.H. 1992. Characterization of a > 900,000-M(r) Cryptosporidium parvum sporozoite glycoprotein recognized by protective hyperimmune bovine colostral immunoglobulin. Infect. Immun. 60, 5132–5138. Petersen, C., Barnes, D.A., and Gousset, L. 1997. Cryptosporidium parvum GP900, a unique invasion protein. J. Eukaryot. Microbiol. 44, 89S–90S. Priest, J.W., Mehlert, A., Arrowood, M.J., Riggs, M.W., and Ferguson, M.A. 2003. Characterization of a low molecular weight glycolipid antigen from Cryptosporidium parvum. J. Biol. Chem. 278, 52212–52222. Ranucci, L., Muller, H.M., La Rosa, G., Reckmann, I., Morales, M.A., Spano, F., Pozio, E., and Crisanti, A. 1993. Characterization and immunolocalization of a Cryptosporidium protein containing repeated amino acid motifs. Infect. Immun. 61, 2347–2356. Rider, S.D., Jr., Cai, X., Sullivan, W.J., Jr., Smith, A.T., Radke, J., White, M., and Zhu, G. 2005. The protozoan parasite Cryptosporidium parvum possesses two functionally and evolutionarily divergent replication protein A large subunits. J. Biol. Chem. 280, 31460–31469. 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. Roberts, C.W., Roberts, F., Henriquez, F.L., Akiyoshi, D., Samuel, B.U., Richards, T.A., Milhous, W., Kyle, D., McIntosh, L., Hill, G.C., Chaudhuri, M., Tzipori, S., and McLeod, R. 2004. Evidence for mitochondrial-derived alternative oxidase in the apicomplexan parasite Cryptosporidium parvum: A potential antimicrobial agent target. Int. J. Parasitol. 34, 297–308. Rochelle, P.A., Fallar, D., Marshall, M.M., Montelone, B.A., Upton, S.J., and Woods, K. 2004. Irreversible UV inactivation of Cryptosporidium spp. despite the presence of UV repair genes. J. Eukaryot. Microbiol. 51, 553–562. Rochelle, P.A., Upton, S.J., Montelone, B.A., and Woods, K. 2005. The response of Cryptosporidium parvum to UV light. Trends Parasitol. 21, 81–87. Rotte, C., Stejskal, F., Zhu, G., Keithly, J.S., and Martin, W. 2001. Pyruvate : NADP+ oxidoreductase from the mitochondrion of Euglena gracilis and from the apicomplexan Cryptosporidium parvum: A biochemical relic linking pyruvate metabolism in mitochondriate and amitochondriate protists. Mol. Biol. Evol. 18, 710–720. Schmatz, D.M. 1989. The mannitol cycle—A new metabolic pathway in the Coccidia. Parasitol. Today 5, 205–208. Senkovich, O., Speed, H., Grigorian, A., Bradley, K., Ramarao, C.S., Lane, B., Zhu, G., and Chattopadhyay, D. 2005. Crystallization of three key glycolytic enzymes of the opportunistic pathogen Cryptosporidium parvum. Biochim. Biophys. Acta 1750, 166–172. Sibley, L.D. 2004. Intracellular parasite invasion strategies. Science 304, 248–253. Slapeta, J. and Keithly, J.S. 2004. Cryptosporidium parvum mitochondrial-type HSP70 targets homologous and heterologous mitochondria. Eukaryot. Cell 3, 483–494. Smith, S. 1994. The animal fatty acid synthase, one gene, one polypeptide, seven enzymes. FASEB J. 8, 1248–1259. Spano, F., Puri, C., Ranucci, L., Putignani, L., and Crisanti, A. 1997. Cloning of the entire COWP gene of Cryptosporidium parvum and ultrastructural localization of the protein during sexual parasite development. Parasitology 114, 427–437. Stein, B., Stover, L., Gillem, A., Winters, K., Leet, J.H., and Chauret, C. 2006. The effect of lectins on Cryptosporidium parvum oocyst in vitro attachment to host cells. J. Parasitol. 92, 1–9. Striepen, B., White, M.W., Li, C., Guerini, M.N., Malik, S.B., Logsdon, J.M., Jr., Liu, C., and Abrahamsen, M.S. 2002. Genetic complementation in apicomplexan parasites. Proc. Natl. Acad. Sci. U.S.A. 99, 6304–6309.


76


Striepen, B. and Kissinger, J.C. 2004. Genomics meets transgenics in search of the elusive Cryptosporidium drug target. Trends Parasitol. 20, 355–358. Strong, W.B. and Nelson, R.G. 2000a. Preliminary profile of the Cryptosporidium parvum genome: An expressed sequence tag and genome survey sequence analysis. Mol. Biochem. Parasitol. 107, 1–32. Strong, W.B. and Nelson, R.G. 2000b. Gene discovery in Cryptosporidium parvum: Expressed sequence tags and genome survey sequences. Contrib. Microbiol. 6, 92–115. Sultan, A.A., Thathy, V., Frevert, U., Robson, K.J., Crisanti, A., Nussenzweig, V., Nussenzweig, R.S., and Menard, R. 1997. TRAP is necessary for gliding motility and infectivity of Plasmodium sporozoites. Cell 90, 511–522. Suzuki, T., Hashimoto, T., Yabu, Y., Kido, Y., Sakamoto, K., Nihei, C., Hato, M., Suzuki, S., Amano, Y., Nagai, K., Hosokawa, T., Minagawa, N., Ohta, N., and Kita, K. 2004. Direct evidence for cyanide-insensitive quinol oxidase (alternative oxidase) in apicomplexan parasite Cryptosporidium parvum: Phylogenetic and therapeutic implications. Biochem. Biophys. Res. Commun. 313, 1044–1052. Takemaru, K., Li, F.Q., Ueda, H., and Hirose, S. 1997. Multiprotein bridging factor 1 (MBF1) is an evolutionarily conserved transcriptional coactivator that connects a regulatory factor and TATA element-binding protein. Proc. Natl. Acad. Sci. U. S. A. 94, 7251–7256. Takemaru, K., Harashima, S., Ueda, H., and Hirose, S. 1998. Yeast coactivator MBF1 mediates GCN4dependent transcriptional activation. Mol. Cell. Biol. 18, 4971–4976. Tekos, A., Prodromaki, E., Papadimou, E., Pavlidou, D., Tsambaos, D., and Drainas, D. 2003. Aminoglycosides suppress tRNA processing in human epidermal keratinocytes in vitro. Skin Pharmacol. Appl. Skin Physiol. 16, 252–258. 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. Theodoulou, F.L., Holdsworth, M., and Baker, A. 2006. Peroxisomal ABC transporters. FEBS Lett. 580, 1139–1155. 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. Tilley, M., Upton, S.J., Blagburn, B.L., and Anderson, B.C. 1990. Identification of outer oocyst wall proteins of three Cryptosporidium (Apicomplexa: Cryptosporidiidae) species by 125I surface labeling. Infect. Immun. 58, 252–253. Tilley, M., Upton, S.J., Fayer, R., Barta, J.R., Chrisp, C.E., Freed, P.S., Blagburn, B.L., Anderson, B.C., and Barnard, S.M. 1991. Identification of a 15-kilodalton surface glycoprotein on sporozoites of Cryptosporidium parvum. Infect. Immun. 59, 1002–1007. Tilley, M. and Upton, S.J. 1994. Both CP15 and CP25 are left as trails behind gliding sporozoites of Cryptosporidium parvum (Apicomplexa). FEMS Microbiol. Lett. 120, 275–278. Umejiego, N.N., Li, C., Riera, T., Hedstrom, L., and Striepen, B. 2004. Cryptosporidium parvum IMP dehydrogenase: Identification of functional, structural, and dynamic properties that can be exploited for drug design. J. Biol. Chem. 279, 40320–40327. Vasquez, J.R., Gooze, L., Kim, K., Gut, J., Petersen, C., and Nelson, R.G. 1996. Potential antifolate resistance determinants and genotypic variation in the bifunctional dihydrofolate reductase-thymidylate synthase gene from human and bovine isolates of Cryptosporidium parvum. Mol. Biochem. Parasitol. 79, 153–165. Voss, T.S., Mini, T., Jenoe, P., and Beck, H.P. 2002. Plasmodium falciparum possesses a cell cycle-regulated short type replication protein A large subunit encoded by an unusual transcript. J. Biol. Chem. 277, 17493–17501. Wanyiri, J.W., O’Connor, R., Allison, G., Kim, K., Kane, A., Qiu, J., Plaut, A.G., and Ward, H.D. 2006. Proteolytic processing of the Cryptosporidium spp. glycoprotein gp40/15 by human furin and by a parasite-derived furin-like protease activity. Infect. Immun. 75, 184–192. Wengelnik, K., Spaccapelo, R., Naitza, S., Robson, K.J., Janse, C.J., Bistoni, F., Waters, A.P., and Crisanti, A. 1999. The A-domain and the thrombospondin-related motif of Plasmodium falciparum TRAP are implicated in the invasion process of mosquito salivary glands. Embo J. 18, 5195–5204.


Biochemistry

77

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.



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

79


80

I.


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,


Epidemiology

81

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


82


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,


Epidemiology

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.


84


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


Epidemiology

85

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


86


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.


Epidemiology

87

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.


88


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


Epidemiology

89

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.


90


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


Epidemiology

91

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).


92


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


Epidemiology

93

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


94


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.


Epidemiology

95

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


96


(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%


Epidemiology

97

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.


98


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).


Epidemiology

99

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.


100


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.)


Epidemiology

101

References Abe, N., Matsubayashi, M., Kimata, I., and Iseki, M. 2006. Subgenotype analysis of Cryptosporidium parvum isolates from humans and animals in Japan using the 60-kDa glycoprotein gene sequences. Parasitol. Res. 99, 303–305. Abe, N., Sawano, Y., Yamada, K., Kimata, I., and Iseki, M. 2002. Cryptosporidium infection in dogs in Osaka, Japan. Vet. Parasitol. 108, 185–193. Adak, G.K., Long, S.M., and O’Brien, S.J. 2002. Trends in indigenous foodborne disease and deaths, England and Wales, 1992 to 2000. Gut. 51, 832–841. Addiss, D.G., Pond, R.S., Remshak, M., Juranek, D.D., Stokes, S., and Davis, J.P. 1996. Reduction of risk of watery diarrhea with point-of-use water filters during a massive outbreak of waterborne Cryptosporidium infection in Milwaukee, Wisconsin, 1993. Am. J. Trop. Med. Hyg. 54, 549–553. Adegbola, R.A., Demba, E., De Veer, G., and Todd, J. 1994. Cryptosporidium infection in Gambian children less than 5 years of age. J. Trop. Med. Hyg. 97, 103–107. Aiello, A.E., Xiao, L., Limor, J.R., Liu, C., Abrahamsen, M.S., and Lal, A.A. 1999. Microsatellite analysis of the human and bovine genotypes of Cryptosporidium parvum. J. Eukaryot. Microbiol. 46, 46S–47S. 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. Alcantara, C.S., Yang, C.H., Steiner, T.S., Barrett, L.J., Lima, A.A., Chappell, C.L., Okhuysen, P.C., White, Jr. A.C., and Guerrant, R.L. 2003. Interleukin-8, tumor necrosis factor-alpha, and lactoferrin in immunocompetent hosts with experimental and Brazilian children with acquired cryptosporidiosis. Am. J. Trop. Med. Hyg. 68, 325–328. Alves, M., Xiao, L., Antunes, F., and Matos, O. 2006. Distribution of Cryptosporidium subtypes in humans and domestic and wild ruminants in Portugal. Parasitol. Res. 99, 287–292. Anon. 1996a. Foodborne outbreak of diarrheal illness associated with Cryptosporidium parvum—Minnesota, 1995. Morb. Mort. Wkly. Rep. 45, 783–784. Anon. 1996b. Outbreak of cryptosporidiosis at a day camp—Florida, July–August 1995. Morb. Mort. Wkly. Rep. 45, 442–444. Anon. 1998. Foodborne outbreak of cryptosporidiosis—Spokane, Washington, 1997. Morb. Mort. Wkly. Rep. 47, 565–567. Anon. 2000. Review of outbreaks of cryptosporidiosis in swimming pools and advice on proceedings of strategic workshop on viability tests and genetic typing. Final report to DEFRA, Drinking Water Inspectorate. Foundation for Water Research, Marlow, Bucks, U.K. (http://www.fwr.org/). Anon. 2001. From the Centers for Disease Control and Prevention. Protracted outbreaks of cryptosporidiosis associated with swimming pool use—Ohio and Nebraska, 2000. JAMA. 285:2967–2969. 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/). Ashbolt, R.H., Coleman, D.J., Misrachi, A., Conti, J.M., and Kirk, M.D. 2003. An outbreak of cryptosporidiosis associated with an animal nursery at a regional fair. Commun. Dis. Intell. 27, 244–249. Atherton, F., Newman, C.P., and Casemore, D.P. 1995. An outbreak of waterborne cryptosporidiosis associated with a public water supply in the U.K. Epidemiol. Infect. 115, 123–131. Badenoch, J. 1990. Cryptosporidium in water supplies. Report of the group of experts. Department of the Environment, Department of Health. London. Badenoch, J. 1995. Cryptosporidium in water supplies. Second report of the group of experts. Department of the Environment, Department of Health. London. Balatbat, A.B., Jordan, G.W., Tang, Y.J., and Silva Jr., J. 1996. Detection of Cryptosporidium parvum DNA in human feces by nested PCR. J. Clin. Microbiol. 34, 1769–1772. Barker, I.K. and Carbonell, P.L. 1974. Cryptosporidium agni sp.n. from lambs, and Cryptosporidium bovis sp.n. from a calf, with observations on the oocyst. Z. Parasitenkd. 44, 289–298. Barwick, R.S., Levy, D.A., Craun, G.F., Beach, M.J., and Calderon, R.L. 2000. Surveillance for waterbornedisease outbreaks—United States, 1997–1998. MMWR CDC Surveill Summ. 49, 1–21. Baxby, D., Hart, C.A., and Taylor. C. 1983. Human cryptosporidiosis, a possible case of hospital cross infection. Br. Med. J. (Clin. Res. Ed). 287, 1760 and 1761.


102


Bell, A., Guasparini, R., Meeds, D., Mathias, R.G., and Farley, J.D. 1993. A swimming pool-associated outbreak of Cryptosporidiosis in British Columbia. Can. J. Pub. Health. 84, 334–337. Bier, J.W. 1991. Isolation of parasites on fruits and vegetables. Southeast Asian J. Trop. Med. Pub. Health. 22 Suppl., 144–145. Black, R.E. 1986. Pathogens that cause travelers’ diarrhea in Latin America and Africa. Rev. Infect. Dis. 8 (Suppl. 2), S131–S135. Blackburn, B.G., Craun, G.F., Yoder, J.S., Hill, V., Calderon, R.L., Chen, N., Lee, S.H. Levy, D.A., and Beach, M.J. 2004. Surveillance for waterborne-disease outbreaks associated with drinking water—United States, 2001–2002. MMWR Surveill Summ. 53, 23–45. Blackburn, B.G., Mazurek, J.M., Hlavsa, M., Park, J., Tillapaw, M., Parrish, M., Salehi, E., Franks, W., Koch, E., Smith, F., Xiao, L., Arrowood, M., Hill, V., da Silva, A., Johnston, S., and Jones, J.L. 2006. Cryptosporidiosis associated with ozonated apple cider. Emerg. Infect. Dis. 12, 684–686. Blagburn, B.L. and Current, W.L. 1983. Accidental infection of a researcher with human Cryptosporidium. J. Infect. Dis. 148, 772 and 773. Bouchier, I. 1998. Cryptosporidium in water supplies; Third report of the group of experts. Department of the Environment, Transport and the Regions, Department of Health. Bridgman, S.A., Robertson, R.M., Syed, Q., Speed, N., Andrews, N., and Hunter, P.R. 1995. Outbreak of cryptosporidiosis associated with a disinfected groundwater supply. Epidemiol. Infect. 115, 555 and 566. Brookes, J.D., Hipsey, M.R., Burch, M.D., Regel, R.H., Linden, L.G., Ferguson, C.M., and Antenucci, J.P. 2005. Relative value of surrogate indicators for detecting pathogens in lakes and reservoirs. Environ. Sci. Technol. 39, 8614–8621. Caccio, S., Homan, W., Camilli, R., Traldi, G., Kortbeek, T., and Pozio, E. 2000. A microsatellite marker reveals population heterogeneity within human and animal genotypes of Cryptosporidium parvum. Parasitology. 120 (Pt. 3), 237–244. Caccio, S., Pinter, E., Fantini, R., Mezzaroma, I., and Pozio, E. 2002. Human infection with Cryptosporidium felis, case report and literature review. Emerg. Infect. Dis. 8, 85 and 86. Caccio, S. and Pozio, E. 2001. Molecular identification of food-borne and water-borne protozoa. Southeast Asian J. Trop. Med. Pub. Health. 32 (Suppl. 2), 156–158. Caccio, S.M. 2005. Molecular epidemiology of human cryptosporidiosis. Parassitologia. 47, 185–192. Calderon, R.L. and Craun, G.F. 2006. Estimates of endemic waterborne risks from community-intervention studies. J. Water Health. 4 (Suppl. 2), 89–99. Cama, V.A., Bern, C., Sulaiman, I.M., Gilman, R.H., Ticona, E., Vivar, A., Kawai, V., Vargas, D., Zhou, L., and Xiao, L. 2003. Cryptosporidium species and genotypes in HIV-positive patients in Lima, Peru. J. Eukaryot. Microbiol. 50 Suppl., 531–533. Campbell, P.N. and Current, W.L. 1983. Demonstration of serum antibodies to Cryptosporidium sp. in normal and immunodeficient humans with confirmed infections. J. Clin. Microbiol. 18, 165–169. Cartwright, R.Y. 2003. Food and waterborne infections associated with package holidays. J. Appl. Microbiol. 94 Suppl., 12S–24S. Casemore, D. 2006. Towards a U.S. national estimate of the risk of endemic waterborne disease—seroepidemiologic studies. J. Water Health. 4 (Suppl. 2), 121–163. Casemore, D.P. 1987. The antibody response to Cryptosporidium, development of a serological test and its use in a study of immunologically normal persons. J. Infect. 14, 125–134. Causer, L.M., Handzel, T., Welch, P., Carr, M., Culp, D., Lucht, R., Mudahar, K., Robinson, D., Neavear, E., Fenton, S., Rose, C., Craig, L., Arrowood, M., Wahlquist, S., Xiao, L., Lee, Y.M., Mirel, L., Levy, D., Beach, M.J., Poquette, G., and Dworkin, M.S. 2006. An outbreak of Cryptosporidium hominis infection at an Illinois recreational waterpark. Epidemiol. Infect. 134, 147–156. Chacin-Bonilla, L. 2001. [Importance of the different species and genotypes of Cryptosporidium in public health]. Invest. Clin. 42, 83–85. Chacin-Bonilla, L. 2002. [Importance of the different species and genotypes of Cryptosporidium in public health]. Invest. Clin. 43, 67–69. Chalmers, R.M., Elwin, K., Thomas, A.L., and Joynson, D.H. 2002a. Infection with unusual types of Cryptosporidium is not restricted to immunocompromised patients. J. Infect. Dis. 185, 270 and 271. Chalmers, R.M., Hughes, S., Thomas, A.L., Woodhouse, S., Thomas, P.D., and Hunter, P. 2002b. Laboratory ascertainment of Cryptosporidium and local authority policies for investigating sporadic cases of cryptosporidiosis in two regions of the United Kingdom. Commun. Dis. Pub. Health. 5, 114–118.


Epidemiology

103

Chalmers, R.M., Sturdee, A.P., Bull, S.A., Miller, A., and Wright, S.E. 1997a. The prevalence of Cryptosporidium parvum and C. muris in Mus domesticus, Apodemus sylvaticus and Clethrionomys glareolus in an agricultural system. Parasitol. Res. 83, 478–482. Chalmers, R.M., Sturdee, A.P., Mellors, P., Nicholson, V., Lawlor, F., Kenny, F., and Timpson, P. 1997b. Cryptosporidium parvum in environmental samples in the Sligo area, Republic of Ireland, a preliminary report. Lett. Appl. Microbiol. 25, 380–384. Chappell, C.L., Okhuysen, P.C., Langer-Curry, R., Widmer, G., Akiyoshi, D.E., Tanriverdi, S., and Tzipori, S. 2006. Cryptosporidium hominis, experimental challenge of healthy adults. Am. J. Trop. Med. Hyg. 75, 851–857. Chappell, C.L., Okhuysen, P.C., Sterling, C.R., and DuPont, H.L. 1996. Cryptosporidium parvum, intensity of infection and oocyst excretion patterns in healthy volunteers. J. Infect. Dis. 173, 232–236. Chappell, C.L., Okhuysen, P.C., Sterling, C.R., Wang, C., Jakubowski, W., and DuPont, H.L. 1999. Infectivity of Cryptosporidium parvum in healthy adults with pre-existing anti-C. parvum serum immunoglobulin G. Am. J. Trop. Med. Hyg. 60, 157–164. Cirak, V.Y. and Bauer, C. 2004. Comparison of conventional coproscopical methods and commercial coproantigen ELISA kits for the detection of Giardia and Cryptosporidium infections in dogs and cats. Berl. Munch. Tierarztl. Wochenschr. 117, 410–413. Clavel, A., Olivares, J.L., Fleta, J., Castillo, J., Varea, M., Ramos, F.J., Arnal, A.C., and Quilez, J. 1996. Seasonality of cryptosporidiosis in children. Eur. J. Clin. Microbiol. Infect. Dis. 15, 77–79. Cohen, S., Dalle, F., Gallay, A., Di Palma, M., Bonnin, A., and Ward, H.D. 2006. Identification of Cpgp40/15 Type Ib as the predominant allele in isolates of Cryptosporidium spp. from a waterborne outbreak of gastroenteritis in South Burgundy, France. J. Clin. Microbiol. 44, 589–591. Colford, J.M., Jr., Roy, S., Beach, M.J., Hightower, A., Shaw, S.E., and Wade, T.J. 2006. A review of household drinking water intervention trials and an approach to the estimation of endemic waterborne gastroenteritis in the United States. J. Wat. Health. 4 (Suppl. 2), 71–88. Collins, M.V., Flick, G.J., Smith, S.A., Fayer, R., Croonenberghs, R., O’Keefe, S., and Lindsay, D.S. 2005a. The effect of high-pressure processing on infectivity of Cryptosporidium parvum oocysts recovered from experimentally exposed Eastern oysters (Crassostrea virginica). J. Eukaryot. Microbiol. 52, 500–504. Collins, M.V., Flick, G.J., Smith, S.A., Fayer, R., Rubendall, E., and Lindsay, D.S. 2005b. The effects of Ebeam irradiation and microwave energy on Eastern Oysters (Crassostrea virginica) experimentally infected with Cryptosporidium parvum. J. Eukaryot. Microbiol. 52, 484–488. Cook, N. 2003. The use of NASBA for the detection of microbial pathogens in food and environmental samples. J. Microbiol. Methods. 53, 165–174. Corso, P.S., Kramer, M.H., Blair, K.A., Addiss, D.G., Davis, J.P., and Haddix, A.C. 2003. Cost of illness in the 1993 waterborne Cryptosporidium outbreak, Milwaukee, Wisconsin. Emerg. Infect. Dis. 9, 426–431. Coupe, S., Sarfati, C., Hamane, S., and Derouin, F. 2005. Detection of Cryptosporidium and identification to the species level by nested PCR and restriction fragment length polymorphism. J. Clin. Microbiol. 43, 1017–1023. Craun, G.F. and Calderon, R.L. 2006a. Observational epidemiologic studies of endemic waterborne risks, cohort, case-control, time-series, and ecologic studies. J. Wat. Health. 4 (Suppl. 2), 101–119. Craun, G.F. and Calderon, R.L. 2006b. Workshop summary, estimating waterborne disease risks in the United States. J. Wat. Health. 4 (Suppl. 2), 241–253. Craun, G.F., Calderon, R.L., and Wade, T.J. 2006a. Assessing waterborne risks: An introduction. J. Wat. Health. 4 (Suppl. 2), 3–18. Craun, M.F., Craun, G.F., Calderon, R.L., and Beach, M.J. 2006b. Waterborne outbreaks reported in the United States. J. Wat. Health. 4 (Suppl. 2), 19–30. Craven, D.E., Steger, K.A., and Hirschhorn, L.R. 1996. Nosocomial colonization and infection in persons infected with human immunodeficiency virus. Infect. Control Hosp. Epidemiol. 17, 304–318. Crawford, F.G. and Vermund, S.H. 1988. Human cryptosporidiosis. Crit. Rev. Microbiol. 16, 113–159. Cruickshank, R., Ashdown, L., and Croese, J. 1988. Human cryptosporidiosis in North Queensland. Aust. N. Z. J. Med. 18, 582–586. Current, W.L., Reese, N.C., Ernst, J.V., Bailey, W.S., Heyman, M.B., and Weinstein, W.M. 1983. Human cryptosporidiosis in immunocompetent and immunodeficient persons: Studies of an outbreak and experimental transmission. N. Engl. J. Med. 308, 1252–1257.


104


Curriero, F.C., Patz, J.A., Rose, J.B., and Lele, S. 2001. The association between extreme precipitation and waterborne disease outbreaks in the United States, 1948–1994. Am. J. Pub. Health. 91, 1194–1199. D’Antonio, R.G., Winn, R.E., Taylor, J.P., Gustafson, T.L., Current, W.L., Rhodes, M.M., Gary, Jr. G.W., and Zajac, R.A. 1985. A waterborne outbreak of cryptosporidiosis in normal hosts. Ann. Intern. Med. 103, 886–888. Dalle, F., Roz, P., Dautin, G., Di Palma, M., Kohli, E., Sire-Bidault, C., Fleischmann, M.G., Gallay, A., Carbonel, S., Bon, F., Tillier, C., Beaudeau, P., and Bonnin, A. 2003. Molecular characterization of isolates of waterborne Cryptosporidium spp. collected during an outbreak of gastroenteritis in South Burgundy, France. J. Clin. Microbiol. 41, 2690–2693. Dann, S.M., Okhuysen, P.C., Salameh, B.M., DuPont, H.L., and Chappell, C.L. 2000. Fecal antibodies to Cryptosporidium parvum in healthy volunteers. Infect. Immun. 68, 5068–5074. Dawson, A., Griffin, R., Fleetwood, A., and Barrett, N.J. 1995. Farm visits and zoonoses. Commun. Dis. Rep. CDR Rev. 5, R81–R86. Dawson, D. 2005. Foodborne protozoan parasites. Int. J. Food Microbiol. 103, 207–227. Dawson, D.J., Samuel, C.M., Scrannage, V., and Atherton, C.J. 2004. Survival of Cryptosporidium species in environments relevant to foods and beverages. J. Appl. Microbiol. 96, 1222–1229. Deng, M.Q. and Cliver, D.O. 2000. Comparative detection of Cryptosporidium parvum oocysts from apple juice. Int. J. Food Microbiol. 54, 155–162. Deng, M.Q., Lam, K.M., and Cliver, D.O. 2000. Immunomagnetic separation of Cryptosporidium parvum oocysts using MACS MicroBeads and high gradient separation columns. J. Microbiol. Meth. 40, 11–17. Dietz, V., Vugia, D., Nelson, R., Wicklund, J., Nadle, J., McCombs, K.G., and Reddy, S. 2000. Active, multisite, laboratory-based surveillance for Cryptosporidium parvum. Am. J. Trop. Med. Hyg. 62, 368–372. DiGiorgio, C.L., Gonzalez, D.A., and Huitt, C.C. 2002. Cryptosporidium and Giardia recoveries in natural waters by using environmental protection agency method 1623. Appl. Environ. Microbiol. 68, 5952–5955. Dillingham, R.A., Lima, A.A., and Guerrant, R.L. 2002. Cryptosporidiosis: Epidemiology and impact. Microbes Infect. 4, 1059–1066. Djuretic, T., Wall, P.G., and Nichols, G. 1997. General outbreaks of infectious intestinal disease associated with milk and dairy products in England and Wales: 1992 to 1996. Commun. Dis. Rep. CDR Rev. 7, R41–R45. Doganci, T., Araz, E., Ensari, A., Tanyuksel, M., and Doganci, L. 2002. Detection of Cryptosporidium parvum infection in childhood using various techniques. Med. Sci. Monit. 8, MT223–MT226. Doria, M.F., Abubakar, I., Syed, Q., Hughes, S., and Hunter, P.R. 2006. Perceived causes of sporadic cryptosporidiosis and their relation to sources of information. J. Epidemiol. Commun. Health. 60, 745–750. Duffy, G. and Moriarty, E.M. 2003. Cryptosporidium and its potential as a food-borne pathogen. Anim. Health Res. Rev. 4, 95–107. Duke, L.A., Breathnach, A.S., Jenkins, D.R., Harkis, B.A., and Codd, A.W. 1996. A mixed outbreak of Cryptosporidium and campylobacter infection associated with a private water supply. Epidemiol. Infect. 116, 303–308. DuPont, H.L., Chappell, C.L., Sterling, C.R., Okhuysen, P.C., Rose, J.B., and Jakubowski, W. 1995. The infectivity of Cryptosporidium parvum in healthy volunteers. N. Engl. J. Med. 332, 855–859. Dworkin, M.S., Goldman, D.P., Wells, T.G., Kobayashi, J.M., and Herwaldt, B.L. 1996. Cryptosporidiosis in Washington State, an outbreak associated with well water. J. Infect. Dis. 174, 1372–1376. Ederli, B.B., Rodrigues, M.F., and Carvalho, C.B. 2005. [Oocysts of the genus Cryptosporidium in domiciliated dogs from the city of Campos dos Goytacazes, the State of Rio de Janeiro]. Rev. Bras. Parasitol.Vet. 14, 129–131. el Sibaei, M.M., Rifaat, M.M., Hameed, D.M., and el Din, H.M. 2003. Nosocomial sources of cryptosporidial infection in newly admitted patients in Ain Shams University Pediatric Hospital. J. Egypt. Soc. Parasitol. 33, 177–188. Elwin, K., Chalmers, R.M., Roberts, R., Guy, E.C., and Casemore, D.P. 2001. Modification of a rapid method for the identification of gene-specific polymorphisms in Cryptosporidium parvum and its application to clinical and epidemiological investigations. Appl. Environ. Microbiol. 67, 5581–5584. Enemark, H.L., Ahrens, P., Juel, C.D., Petersen, E., Petersen, R.F., Andersen, J.S., Lind, P., and Thamsborg, S.M. 2002. Molecular characterization of Danish Cryptosporidium parvum isolates. Parasitology. 125, 331–341.


Epidemiology

105

Esteban, J.G., Aguirre, C., Flores, A., Strauss, W., Angles, R., and Mas-Coma, S. 1998. High Cryptosporidium prevalences in healthy Aymara children from the northern Bolivian Altiplano. Am. J. Trop. Med. Hyg. 58, 50–55. Evans, M.R. and Gardner, D. 1996. Cryptosporidiosis outbreak associated with an educational farm holiday. Commun. Dis. Rep. CDR Rev. 6, R50 and R51. Exner, M. and Gornik, V. 2004. [Parasitic zoonoses transmitted by drinking water. Giardiasis and cryptosporidiosis]. Bundesgesundheitsblatt. Gesundheitsforschung. Gesundheitsschutz. 47, 698–704. Fayer, R., Lewis, E.J., Trout, J.M., Graczyk, T.K., Jenkins, M.C., Higgins, J., Xiao, L., and Lal, A.A. 1999. Cryptosporidium parvum in oysters from commercial harvesting sites in the Chesapeake Bay. Emerg. Infect. Dis. 5, 706–710. Fayer, R., Trout, J.M., Lewis, E.J., Xiao, L., Lal, A., Jenkins, M.C., and Graczyk, T.K. 2002. Temporal variability of Cryptosporidium in the Chesapeake Bay. Parasitol. Res. 88, 998–1003. Fayer, R. and Ungar, B.L. 1986. Cryptosporidium spp. and cryptosporidiosis. Microbiol. Rev. 50, 458–483. Flint, J.A., Van Duynhoven, Y.T., Angulo, F.J., DeLong, S.M., Braun, P., Kirk, M., Scallan, E., Fitzgerald, M., Adak, G.K., Sockett, P., Ellis, A., Hall, G., Gargouri, N., Walke, H., and Braam, P. 2005. Estimating the burden of acute gastroenteritis, foodborne disease, and pathogens commonly transmitted by food: An international review. Clin. Infect. Dis. 41, 698–704. Fontanarrosa, M.F., Vezzani, D., Basabe, J., and Eiras, D.F. 2006. An epidemiological study of gastrointestinal parasites of dogs from Southern Greater Buenos Aires (Argentina), age, gender, breed, mixed infections, and seasonal and spatial patterns. Vet. Parasitol. 136, 283–295. Franco, R.M. and Cantusio, N.R. 2002. Occurrence of cryptosporidial oocysts and Giardia cysts in bottled mineral water commercialized in the city of Campinas, State of Sao Paulo, Brazil. Mem. Inst. Oswaldo Cruz. 97, 205–207. Freire-Santos, F., Oteiza-Lopez, A.M., Castro-Hermida, J.A., Garcia-Martin, O., and Ares-Mazas, M.E. 2001. Viability and infectivity of oocysts recovered from clams, Ruditapes philippinarum, experimentally contaminated with Cryptosporidium parvum. Parasitol. Res. 87, 428–430. Fripp, P.J., Bothma, M.T., and Crewe-Brown, H.H. 1991. Four years of cryptosporidiosis at GaRankuwa Hospital. J. Infect. 23, 93–100. Frost, F., Craun, G., Mihaly, K., Gyorgy, B., Calderon, R., and Muller, T. 2005a. Serological responses to Cryptosporidium antigens among women using riverbank-filtered water, conventionally filtered surface water and groundwater in Hungary. J. Wat. Health. 3, 77–82. Frost, F.J., Calderon, R.L., Muller, T.B., Curry, M., Rodman, J.S., Moss, D.M., and de la Cruz, A.A. 1998. A two-year follow-up survey of antibody to Cryptosporidium in Jackson County, Oregon following an outbreak of waterborne disease. Epidemiol. Infect. 121, 213–217. Frost, F.J., Kunde, T.R., Muller, T.B., Craun, G.F., Katz, L.M., Hibbard, A.J., and Calderon, R.L. 2003. Serological responses to Cryptosporidium antigens among users of surface- vs. ground-water sources. Epidemiol. Infect. 131, 1131–1138. Frost, F.J., Muller, T., Craun, G.F., Fraser, D., Thompson, D., Notenboom, R., and Calderon. R.L. 2000. Serological analysis of a cryptosporidiosis epidemic. Int. Epidemiol. 29, 376–379. Frost, F.J., Muller, T., Craun, G.F., Lockwood, W.B., and Calderon, R.L. 2002. Serological evidence of endemic waterborne Cryptosporidium infections. Ann. Epidemiol. 12, 222–227. Frost, F.J., Roberts, M., Kunde, T.R., Craun, G., Tollestrup, K., Harter, L., and Muller, T. 2005b. How clean must our drinking water be: The importance of protective immunity. J. Infect. Dis. 191, 809–814. Frost, F.J., Tollestrup, K., Craun, G.F., Fairley, C.K., Sinclair, M.I., and Kunde, T.R. 2005c. Protective immunity associated with a strong serological response to a Cryptosporidium-specific antigen group, in HIV-infected individuals. J. Infect. Dis. 192, 618–621. Furtado, C., Adak, G.K., Stuart, J.M., Wall, P.G., Evans, H.S., and Casemore, D.P. 1998. Outbreaks of waterborne infectious intestinal disease in England and Wales, 1992–5. Epidemiol. Infect. 121, 109–119. Gale, P. 1996. Developments in microbiological risk assessment models for drinking water—a short review. J. Appl. Bacteriol. 81, 403–410. Gale, P. 2003. Using event trees to quantify pathogen levels on root crops from land application of treated sewage sludge. J. Appl. Microbiol. 94, 35–47. Gale, P. 2005. Land application of treated sewage sludge: Quantifying pathogen risks from consumption of crops. J. Appl. Microbiol. 98, 380–396.


106


Garcia, L., Henderson, J., Fabri, M., and Oke, M. 2006. Potential sources of microbial contamination in unpasteurized apple cider. J. Food Prot. 69, 137–144. Gardner, C. 1994. An outbreak of hospital-acquired cryptosporidiosis. Br. J. Nurs. 3, 152, 154–158. Gatei, W., Ashford, R.W., Beeching, N.J., Kamwati, S.K., Greensill, J., and Hart, C.A. 2002a. Cryptosporidium muris infection in an HIV-infected adult, Kenya. Emerg. Infect. Dis. 8, 204–206. Gatei, W., Das, P., Dutta, P., Sen, A., Cama, V., Lal, A.A., and Xiao, L. 2007. Multilocus sequence typing and genetic structure of Cryptosporidium hominis from children in Kolkata, India. Infect. Genet. Evol. 7, 197–205. Gatei, W., Greensill, J., Ashford, R.W., Cuevas, L.E., Parry, C.M., Cunliffe, N.A., Beeching, N.J., and Hart, C.A. 2003. Molecular analysis of the 18S rRNA gene of Cryptosporidium parasites from patients with or without human immunodeficiency virus infections living in Kenya, Malawi, Brazil, the United Kingdom, and Vietnam. J. Clin. Microbiol. 41, 1458–1462. Gatei, W., Suputtamongkol, Y., Waywa, D., Ashford, R.W., Bailey, J.W., Greensill, J., Beeching, N.J., and Hart, C.A. 2002b. Zoonotic species of Cryptosporidium are as prevalent as the anthroponotic in HIVinfected patients in Thailand. Ann. Trop. Med. Parasitol. 96, 797–802. Gelletlie, R., Stuart, J., Soltanpoor, N., Armstrong, R., and Nichols, G. 1997. Cryptosporidiosis associated with school milk. Lancet. 350, 1005 and 1006. Glaberman, S., Moore, J.E., Lowery, C.J., Chalmers, R.M., Sulaiman, I., Elwin, K., Rooney, P.J., Millar, B.C., Dooley, J.S., Lal, A.A., and Xiao, L. 2002. Three drinking-water-associated cryptosporidiosis outbreaks, Northern Ireland. Emerg. Infect. Dis. 8, 631–633. Glaberman, S., Sulaiman, I.M., Bern, C., J., Limor, Peng, M.M., Morgan, U., Gilman, R., Lal, A.A., and Xiao, L. 2001. A multilocus genotypic analysis of Cryptosporidium meleagridis. J. Eukaryot. Microbiol. Suppl., 19S–22S. Gobet, P. and Toze, S. 2001. Sensitive genotyping of Cryptosporidium parvum by PCR-RFLP analysis of the 70-kilodalton heat shock protein (HSP70) gene. FEMS Microbiol. Lett. 200, 37–41. Goh, S., Reacher, M., Casemore, D.P., Verlander, N.Q., Chalmers, R., Knowles, M., Williams, J., Osborn, K., and Richards, S. 2004. Sporadic cryptosporidiosis, North Cumbria, England, 1996–2000. Emerg. Infect. Dis. 10, 1007–1015. Goh, S., Reacher, M., Casemore, D.P., Verlander, N.Q., Charlett, A., Chalmers, R.M., Knowles, M., Pennington, A., Williams, J., Osborn, K., and Richards, S. 2005. Sporadic cryptosporidiosis decline after membrane filtration of public water supplies, England, 1996–2002. Emerg. Infect. Dis. 11, 251–259. Goldstein, S.T., Juranek, D.D., Ravenholt, O., Hightower, A.W., Martin, D.G., Mesnik, J.L., Griffiths, S.D., Bryant, A.J., Reich, R.R., and Herwaldt, B.L. 1996. Cryptosporidiosis: An outbreak associated with drinking water despite state-of-the-art water treatment. Ann. Intern. Med. 124, 459–468. Gomez-Couso, H., Mendez-Hermida, F., Castro-Hermida, J.A., and Ares-Mazas, E. 2006. Cryptosporidium contamination in harvesting areas of bivalve molluscs. J. Food Prot. 69, 185–190. Graczyk, T.K., Cranfield, M.R., Fayer, R., and Bixler, H. 1999a. House flies (Musca domestica) as transport hosts of Cryptosporidium parvum. Am. J. Trop. Med. Hyg. 61, 500–504. Graczyk, T.K., Fayer, R., Cranfield, M.R., Mhangami-Ruwende, B., Knight, R., Trout, J.M., and Bixler, H. 1999b. Filth flies are transport hosts of Cryptosporidium parvum. Emerg. Infect. Dis. 5, 726 and 727. Graczyk, T.K., Fayer, R., Knight, R., Mhangami-Ruwende, B., Trout, J.M., da Silva, A.J., and Pieniazek, N.J. 2000. Mechanical transport and transmission of Cryptosporidium parvum oocysts by wild filth flies. Am. J. Trop. Med. Hyg. 63, 178–183. Graczyk, T.K., Fayer, R., Lewis, E.J., Trout, J.M., and Farley, C.A. 1999c. Cryptosporidium oocysts in Bent mussels (Ischadium recurvum) in the Chesapeake Bay. Parasitol. Res. 85, 518–521. Graczyk, T.K., Marcogliese, D.J., De Lafontaine, Y., da Silva, A.J., Mhangami-Ruwende, B., and Pieniazek, N.J. 2001 Cryptosporidium parvum oocysts in zebra mussels (Dreissena polymorpha), evidence from the St Lawrence River. Parasitol. Res. 87, 231–234. Graczyk, T.K. and Schwab, K.J. 2000. Foodborne infections vectored by molluscan shellfish. Curr. Gastroenterol. Rep. 2, 305–309. Gutteridge, W. and Haworth, E.A. 1994. An outbreak of gastrointestinal illness associated with contamination of the mains supply by river water. Commun Dis. Rep. CDR Review. 4, R50–51. Guyot, K., Follet-Dumoulin, A., Lelievre, E., Sarfati, C., Rabodonirina, M., Nevez, G., Cailliez, J.C., Camus, D., and Dei-Cas, E. 2001. Molecular characterization of Cryptosporidium isolates obtained from humans in France. J. Clin. Microbiol. 39, 3472–3480.


Epidemiology

107

Guyot, K., Follet-Dumoulin, A., Recourt, C., Lelievre, E., Cailliez, J.C., and Dei-Cas, E. 2002. PCR-restriction fragment length polymorphism analysis of a diagnostic 452-base-pair DNA fragment discriminates between Cryptosporidium parvum and C. meleagridis and between C. parvum isolates of human and animal origin. Appl. Environ. Microbiol. 68, 2071–2076. Hackett, T. and Lappin, M.R. 2003. Prevalence of enteric pathogens in dogs of north-central Colorado. J. Am. Anim. Hosp. Assoc. 39, 52–56. Hajdusek, O., Ditrich, O., and Slapeta, J. 2004. Molecular identification of Cryptosporidium spp. in animal and human hosts from the Czech Republic. Vet. Parasitol. 122, 183–192. Hanes, D.E., Worobo, R.W., Orlandi, P.A., Burr, D.H., Miliotis, M.D., Robl, M.G., Bier, J.W., Arrowood, M.J., Churey, J.J., and Jackson, G.I. 2002. Inactivation of Cryptosporidium parvum oocysts in fresh apple cider by UV irradiation. Appl. Environ. Microbiol. 68, 4168–4172. Hannah, J. and Riordan, T. 1988. Case to case spread of cryptosporidiosis; evidence from a day nursery outbreak. Pub. Health. 102, 539–544. Hansen, J.S. and Ongerth, J.E. 1991. Effects of time and watershed characteristics on the concentration of Cryptosporidium oocysts in river water. Appl. Environ. Microbiol. 57, 2790–2795. Harrison, S.L., Nelder, R., Hayek, L., MacKenzie, I.F., Casemore, D.P., and Dance, D. 2002. Managing a large outbreak of cryptosporidiosis, how to investigate and when to decide to lift a “boil water” notice. Commun. Dis. Pub. Health. 5, 230–239. Havelaar, A.H., De Hollander, A.E., Teunis, P.F., Evers, E.G., Van Kranen, H.J., Versteegh, J.F., Van Koten, J.E., and Slob, W. 2000. Balancing the risks and benefits of drinking water disinfection, disability adjusted life-years on the scale. Environ. Health Perspect. 108, 315–321. Hayes, E.B., Matte, T.D., O’Brien, T.R., McKinley, T.W., Logsdon, G.S., Rose, J.B., Ungar, B.L., Word, D.M., Pinsky, P.F., and Cummings, M.L. 1989. Large community outbreak of cryptosporidiosis due to contamination of a filtered public water supply. N. Engl. J. Med. 320, 1372–1376. Hellard, M., Hocking, J., Willis, J., Dore, G., and Fairley, C. 2003. Risk factors leading to Cryptosporidium infection in men who have sex with men. Sex Transm. Infect. 79, 412–414. Hiepe, T. and Buchwalder, R. 1991. [Livestock manure as a vector for parasites—a report of experiences]. Dtsch. Tierarztl. Wochenschr. 98, 268–272. Hlavsa, M.C., Watson, J.C., and Beach, M.J. 2005. Cryptosporidiosis surveillance—United States 1999–2002. MMWR Surveill. Summ. 54, 1–8. Horen, W.P. 1983. Detection of Cryptosporidium in human fecal specimens. J. Parasitol. 69, 622–624. Horman, A., Korpela, H., Sutinen, J., Wedel, H., and Hanninen, M.L. 2004. Meta-analysis in assessment of the prevalence and annual incidence of Giardia spp. and Cryptosporidium spp. infections in humans in the Nordic countries. Int. J. Parasitol. 34, 1337–1346. Howard, G., Pedley, S., and Tibatemwa, S. 2006. Quantitative microbial risk assessment to estimate health risks attributable to water supply, can the technique be applied in developing countries with limited data? J. Water Health. 4, 49–65. Howe, A.D., Forster, S., Morton, S., Marshall, R., Osborn, K.S., Wright, P., and Hunter, P.R. 2002. Cryptosporidium oocysts in a water supply associated with a cryptosporidiosis outbreak. Emerg. Infect. Dis. 8, 619–624. Hoxie, N.J., Davis, J.P., Vergeront, J.M., Nashold, R.D., and Blair, K.A. 1997. Cryptosporidiosis-associated mortality following a massive waterborne outbreak in Milwaukee, Wisconsin. Am. J. Pub. Health. 87, 2032–2035. Hrudy, S.E. and Hrudy, E.J. 2004. Safe Drinking water: Lessons from recent outbreaks in affluent nations. IWA. London. Hsu, B.M. and Yeh, H.H. 2003. Removal of Giardia and Cryptosporidium in drinking water treatment, a pilotscale study. Wat. Res. 37, 1111–1117. Huber, F., Bomfim, T.C., and Gomes, R.S. 2005. Comparison between natural infection by Cryptosporidium sp., Giardia sp. in dogs in two living situations in the West Zone of the municipality of Rio de Janeiro. Vet. Parasitol. 130, 69–72. Hunt, D.A., Sebugwawo, S., Edmondson, S.G., and Casemore, D.P. 1994. Cryptosporidiosis associated with a swimming pool complex. Commun. Dis. Rep. CDR Rev. 4, R20–R22. Hunter, P.R., Chalmers, R.M., Syed, Q., Hughes, L.S., Woodhouse, S., and Swift, L. 2003. Foot and mouth disease and cryptosporidiosis, possible interaction between two emerging infectious diseases. Emerg. Infect. Dis. 9, 109–112.


108


Hunter, P.R., Hughes, S., Woodhouse, S., Raj, N., Syed, Q., Chalmers, R.M., Verlander, N.Q., and Goodacre, J. 2004a. Health sequelae of human cryptosporidiosis in immunocompetent patients. Clin. Infect. Dis. 39, 504–510. Hunter, P.R., Hughes, S., Woodhouse, S., Syed, Q., Verlander, N.Q., Chalmers, R.M., Morgan, K., Nichols, G., Beeching, N., and Osborn, K. 2004b. Sporadic cryptosporidiosis case-control study with genotyping. Emerg. Infect. Dis. 10, 1241–1249. Hunter, P.R. and Nichols, G. 2002. Epidemiology and clinical features of Cryptosporidium infection in immunocompromised patients. Clin. Microbiol. Rev. 15, 145–154. Hunter, P.R. and Syed, Q. 2001. Community surveys of self-reported diarrhea can dramatically overestimate the size of outbreaks of waterborne cryptosporidiosis. Water Sci. Technol. 43, 27–30. Hunter, P.R. and Syed, Q. 2002. A community survey of self-reported gastroenteritis undertaken during an outbreak of cryptosporidiosis strongly associated with drinking water after much press interest. Epidemiol. Infect. 128, 433–438. Insulander, M., Lebbad, M., Stenstrom, T.A., and Svenungsson, B. 2005. An outbreak of cryptosporidiosis associated with exposure to swimming pool water. Scand. J. Infect. Dis. 37, 354–360. Iqbal, J., Hira, P.R., Al Ali, F., and Philip, R. 2001. Cryptosporidiosis in Kuwaiti children, seasonality and endemicity. Clin. Microbiol. Infect. 7, 261–266. Jelinek, T., Lotze, M., Eichenlaub, S., Loscher, T., and Nothdurft, H.D. 1997. Prevalence of infection with Cryptosporidium parvum and Cyclospora cayetanensis among international travellers. Gut. 41, 801–804. Joachim, A. 2004. [Is Cryptosporidium a zoonotic agent?]. Wien. Klin. Wochenschr. 116 (Suppl. 4), 2–6. Joce, R.E., Bruce, J., Kiely, D., Noah, N.D., Dempster, W.B., Stalker, R., Gumsley, P., Chapman, P.A., Norman, P., and Watkins, J. 1991. An outbreak of cryptosporidiosis associated with a swimming pool. Epidemiol. Infect. 107, 497–508. Jokipii, L., Pohjola, S., and Jokipii, A.M. 1985. Cryptosporidiosis associated with traveling and Giardiasis. Gastroenterology. 89, 838–842. Jones, M., Boccia, D., Kealy, M., Salkin, B., Ferrero, A., Nichols, G., and Stuart, J. 2006. Cryptosporidium outbreak linked to interactive water feature, U.K.: importance of guidelines. Euro. Surveill. 11. Joseph, C., Hamilton, G., O’Connor, M., Nicholas, S., Marshall, R., Stanwell-Smith, R., Sims, R., Ndawula, E., Casemore, D., and Gallagher, P. 1991. Cryptosporidiosis in the Isle of Thanet; an outbreak associated with local drinking water. Epidemiol. Infect. 107, 509–519. Jougla, E., Pequignot, F., Carbon, C., Pavillon, G., Mireille, E.B., Bourdais, J.P., Bourdais, O., and Hatton, F. 1996. AIDS-related conditions, study of a representative sample of 1203 patients deceased in 1992 in France. Int. J. Epidemiol. 25, 190–197. Katsumata, T., Hosea, D., Ranuh, I.G., Uga, S., Yanagi, T., and Kohno, S. 2000. Short report: Possible Cryptosporidium muris infection in humans. Am. J. Trop. Med. Hyg. 62, 70–72. Khalakdina, A., Vugia, D.J., Nadle, J., Rothrock, G.A., and Colford, Jr. J.M. 2003. Is drinking water a risk factor for endemic cryptosporidiosis? A case-control study in the immunocompetent general population of the San Francisco Bay Area. BMC. Pub. Health. 3, 11. Kiang, K.M., Scheftel, J.M., Leano, F.T., Taylor, C.M., Belle-Isle, P.A., Cebelinski, E.A., Danila, R., and Smith, K.E. 2006. Recurrent outbreaks of cryptosporidiosis associated with calves among students at an educational farm programme, Minnesota, 2003. Epidemiol. Infect. 134, 878–886. Kim, L.S., Stansell, J., Cello, J.P., and Koch, J. 1998. Discrepancy between sex- and water-associated risk behaviors for cryptosporidiosis among HIV-infected patients in San Francisco. J. Acquir. Immune. Defic. Syndr. Hum. Retrovirol. 19, 44–49. Kjos, S.A., Jenkins, M., Okhuysen, P.C., and Chappell, C.L. 2005. Evaluation of recombinant oocyst protein CP41 for detection of Cryptosporidium-specific antibodies. Clin. Diagn. Lab Immunol. 12, 268–272. Kniel, K.E. and Jenkins, M.C. 2005. Detection of Cryptosporidium parvum oocysts on fresh vegetables and herbs using antibodies specific for a Cryptosporidium parvum viral antigen. J. Food Prot. 68, 1093–1096. Kniel, K.E., Sumner, S.S., Lindsay, D.S., Hackney, C.R., Pierson, M.D., Zajac, A.M., Golden, D.A., and Fayer, R. 2003. Effect of organic acids and hydrogen peroxide on Cryptosporidium parvum viability in fruit juices. J. Food Prot. 66, 1650–1657. Koch, K.L., Phillips, D.J., Aber, R.C., and Current, W.L. 1985. Cryptosporidiosis in hospital personnel: Evidence for person-to-person transmission. Ann. Intern. Med. 102, 593–596.


Epidemiology

109

Kramer, M.H., Sorhage, F.E., Goldstein, S.T., Dalley, E., Wahlquist, S.P., and Herwaldt, B.L. 1998. First reported outbreak in the United States of cryptosporidiosis associated with a recreational lake. Clin. Infect. Dis. 26, 27–33. Krause, W., Abraham, A., and Lehmann, D. 1995. [Evidence of Cryptosporidium in children with symptomatic enteritis from the Leipzig administrative area 1987–1992]. Appl. Parasitol. 36, 66–71. Kuroki, T., Watanabe, Y., Asai, Y., Yamai, S., Endo, T., Uni, S., Kimata, I., and Iseki, M. 1996. [An outbreak of waterborne Cryptosporidiosis in Kanagawa, Japan]. Kansensh. Zasshi. 70, 132–140. Laberge, I., Ibrahim, A., Barta, J.R., and Griffiths, M.W. 1996. Detection of Cryptosporidium parvum in raw milk by PCR and oligonucleotide probe hybridization. Appl. Environ. Microbiol. 62, 3259–3264. Lake, I.R., Bentham, G., Kovats, R.S., and Nichols, G.L. 2005. Effects of weather and river flow on cryptosporidiosis. J. Wat. Health. 3, 469–474. Lake, I.R., Nichols, G., Bentham, G., Harrison, F.C., Hunter, P.R., and Kovats, R.S. 2007. Cryptosporidiosis decline after regulation, England and Wales, 1989–2005. Emerg. Infect. Dis. 13: 623–625. Lallo, M.A. and Bondan, E.F. 2006. [Prevalence of Cryptosporidium sp. in institutionalized dogs in the city of Sao Paulo, Brazil]. Rev. Saude Publica. 40, 120–125. Laupland, K.B. and Church, D.L. 2005. Population-based laboratory surveillance for Giardia sp. and Cryptosporidium sp. infections in a large Canadian health region. BMC. Infect. Dis. 5, 72. Leach, C.T., Koo, F.C., Kuhls, T.L., Hilsenbeck, S.G., and Jenson, H.B. 2000. Prevalence of Cryptosporidium parvum infection in children along the Texas-Mexico border and associated risk factors. Am. J. Trop. Med. Hyg. 62, 656–661. Learmonth, J.J., Ionas, G., Ebbett, K.A., and Kwan, E.S. 2004. Genetic characterization and transmission cycles of Cryptosporidium species isolated from humans in New Zealand. Appl. Environ. Microbiol. 70, 3973–3978. Learmonth, J.J., Ionas, G., Pita, A.B., and Cowie, R.S. 2003. Identification and genetic characterisation of Giardia and Cryptosporidium strains in humans and dairy cattle in the Waikato Region of New Zealand. Wat. Sci.Technol. 47, 21–26. Leav, B.A., Mackay, M.R., Anyanwu, A., O’ Connor, R.M., Cevallos, A.M., Kindra, G., Rollins, N.C., Bennish, M.L., Nelson, R.G., and Ward, H.D. 2002. Analysis of sequence diversity at the highly polymorphic Cpgp40/15 locus among Cryptosporidium isolates from human immunodeficiency virus-infected children in South Africa. Infect. Immun. 70, 3881–3890. Lee, S.H., Levy, D.A., Craun, G.F., Beach, M.J., and Calderon, R.L. 2002. Surveillance for waterborne-disease outbreaks—United States, 1999–2000. MMWR Surveill. Summ. 51, 1–47. Lemmon, J.M., McAnulty, J.M., and Bawden-Smith, J. 1996. Outbreak of cryptosporidiosis linked to an indoor swimming pool. Med. J. Aust. 165, 613–616. Leoni, F., Amar, C., Nichols, G., Pedraza-Diaz, S., and McLauchlin, J. 2006a. Genetic analysis of Cryptosporidium from 2414 humans with diarrhea in England between 1985 and 2000. J. Med. Microbiol. 55, 703–707. Leoni, F., Amar, C., Nichols, G., Pedraza-Diaz, S., and McLauchlin, J. 2006b. Genetic analysis of Cryptosporidium from 2414 humans with diarrhea in England between 1985 and 2000. J. Med. Microbiol. 55, 703–707. 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., Gallimore, C.I. Green, J., and McLauchlin, J. 2006c. Characterisation of small double stranded RNA molecule in Cryptosporidium hominis, Cryptosporidium felis and Cryptosporidium meleagridis. Parasitol. Int. 55, 299–306. Leoni, F., Mallon, M.E., Smith, H.V., Tait, A., and McLauchlin, J. 2007. Multilocus analysis of Cryptosporidium hominis and Cryptosporidium parvum from sporadic and outbreak-related human cases and C. parvum from sporadic cases in livestock in the U.K. J. Clin. Microbiol. Submitted for publication. Li, X., Atwill, E.R., Dunbar, L.A., Jones, T., Hook, J., and Tate, K.W. 2005. Seasonal temperature fluctuations induces rapid inactivation of Cryptosporidium parvum. Environ. Sci. Technol. 39, 4484–4489. Li, X., Guyot, K., Dei-Cas, E., Mallard, J.P., Ballet, J.J., and Brasseur, P. 2006. Cryptosporidium oocysts in mussels (Mytilus edulis) from Normandy (France). Int. J. Food Microbiol. 108, 321–325. Lim, L.S., Varkey, P., Giesen, P., and Edmonson, L. 2004. Cryptosporidiosis outbreak in a recreational swimming pool in Minnesota. J. Environ. Health. 67, 16–20, 28, 27.


110


Lindergard, G., Nydam, D.V., Wade, S.E., Schaaf, S.L., and Mohammed, H.O. 2003. A novel multiplex polymerase chain reaction approach for detection of four human infective Cryptosporidium isolates, Cryptosporidium parvum, types H and C, Cryptosporidium canis, and Cryptosporidium felis in fecal and soil samples. J. Vet. Diagn. Invest. 15, 262–267. Lopez-Velez, R., Tarazona, R., Garcia, C.A., Gomez-Mampaso, E., Guerrero, A., Moreira, V., and Villanueva, R. 1995. Intestinal and extraintestinal cryptosporidiosis in AIDS patients. Eur. J. Clin. Microbiol. Infect. Dis. 14, 677–681. Louie, K., Gustafson, L., Fyfe, M., Gill, I., MacDougall, L., Tom, L., Wong, Q., and Isaac-Renton, J. 2004. An outbreak of Cryptosporidium parvum in a Surrey pool with detection in pool water sampling. Can. Commun. Dis. Rep. 30, 61–66. Ma, P. and Soave, R. 1983. Three-step stool examination for cryptosporidiosis in 10 homosexual men with protracted watery diarrhea. J. Infect. Dis. 147, 824–828. MacKenzie, W.R., Hoxie, N.J., Proctor, M.E., Gradus, M.S., Blair, K.A., Peterson, D.E., Kazmierczak, J.J., Addiss, D.G., Fox, K.R., and Rose, J.B. 1994. A massive outbreak in Milwaukee of Cryptosporidium infection transmitted through the public water supply. N. Engl. J. Med. 331, 161–167. MacKenzie, W.R., Kazmierczak, J.J., and Davis, J.P. 1995. An outbreak of cryptosporidiosis associated with a resort swimming pool. Epidemiol. Infect. 115, 545–553. MacRae, M., Hamilton, C., Strachan, N.J., Wright, S., and Ogden, I.D. 2005. The detection of Cryptosporidium parvum and Escherichia coli O157 in U.K. bivalve shellfish. J. Microbiol. Methods. 60, 395–401. Maguire, H.C., Holmes, E., Hollyer, J., Strangeways, J.E., Foster, P., Holliman, R.E., and Stanwell-Smith, R. 1995. An outbreak of cryptosporidiosis in south London, what value the p value? Epidemiol. Infect. 115, 279–287. 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. Mallon, M., Macleod, A., Wastling, J., Smith, H., Reilly, B., and Tait, A. 2003a. Population structures and the role of genetic exchange in the zoonotic pathogen Cryptosporidium parvum. J. Mol. Evol. 56, 407–417. Mallon, M.E., Macleod, A., Wastling, J.M., Smith, H., and Tait, A. 2003b. Multilocus genotyping of Cryptosporidium parvum Type 2, population genetics and sub-structuring. Infect. Genet. Evol. 3, 207–218. Martino, P., Gentile, G., Caprioli, A., Baldassarri, L., Donelli, G., Arcese, W., Fenu, S., Micozzi, A., Venditti, M., and Mandelli, F. 1988. Hospital-acquired cryptosporidiosis in a bone marrow transplantation unit. J. Infect. Dis. 158, 647–648. Mathieu, E., Levy, D.A., Veverka, F., Parrish, M.K., Sarisky, J., Shapiro, N., Johnston, S., Handzel, T., Hightower, A., Xiao, L., Lee, Y.M., York, S., Arrowood, M., Lee, R., and Jones, J.L. 2004. Epidemiologic and environmental investigation of a recreational water outbreak caused by two genotypes of Cryptosporidium parvum in Ohio in 2000. Am. J. Trop. Med. Hyg. 71, 582–589. 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 and 2257. McAnulty, J.M., Fleming, D.W., and Gonzalez, A.H. 1994. A community-wide outbreak of cryptosporidiosis associated with swimming at a wave pool. J. Am. Med. Assoc. 272, 1597–1600. McAnulty, J.M., Keene, W.E., Leland, D., Hoesly, F., Hinds, B., Stevens, G., and Fleming, D.W. 2000. Contaminated drinking water in one town manifesting as an outbreak of cryptosporidiosis in another. Epidemiol. Infect. 125, 79–86. McDonald, A.C., MacKenzie, W.R., Addiss, D.G., Gradus, M.S., Linke, G., Zembrowski, E., Hurd, M.R., Arrowood, M.J., Lammie, P.J., and Priest, J.W. 2001. Cryptosporidium parvum-specific antibody responses among children residing in Milwaukee during the 1993 waterborne outbreak. J. Infect. Dis. 183, 1373–1379. McGlade, T.R., Robertson, I.D., Elliot, A.D., Read, C., and Thompson, R.C. 2003. Gastrointestinal parasites of domestic cats in Perth, Western Australia. Vet. Parasitol. 117, 251–262. 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. McLauchlin, J., Amar, C., 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, 329–335.


Epidemiology

111

McLauchlin, J., Casemore, D.P., Moran, S., and Patel, S. 1998. The epidemiology of cryptosporidiosis: Application of experimental sub-typing and antibody detection systems to the investigation of waterborne outbreaks. Folia Parasitol. (Praha). 45, 83–92. McLauchlin, J., Pedraza-Diaz, S., Amar-Hoetzeneder, C., and Nichols, G.L. 1999. Genetic characterization of Cryptosporidium strains from 218 patients with diarrhea diagnosed as having sporadic cryptosporidiosis. J. Clin. Microbiol. 37, 3153–3158. Mead, P.S., Slutsker, L., Dietz, V., McCaig, L.F., Bresee, J.S., Shapiro, C., Griffin, P.M., and Tauxe, R.V. 1999. Food-related illness and death in the United States. Emerg. Infect. Dis. 5, 607–625. Medema, G.J., Hoogenboezem, W., van der Veer, A.J., Ketelaars, H.A., Hijnen, W.A., and Nobel, P.J. 2003. Quantitative risk assessment of Cryptosporidium in surface water treatment. Wat. Sci. Technol. 47, 241–247. Meisel, J.L., Perera, D.R., Meligro, C., and Rubin, C.E. 1976. Overwhelming watery diarrhea associated with Cryptosporidium in an immunosuppressed patient. Gastroenterology 70, 1156–1160. Messner, M., Shaw, S., Regli, S., Rotert, K., Blank, V., and Soller, J. 2006. An approach for developing a national estimate of waterborne disease due to drinking water and a national estimate model application. J. Wat. Health. 4 (Suppl. 2), 201–240. Messner, M.J., Chappell, C.L., and Okhuysen, P.C. 2001. Risk assessment for Cryptosporidium: A hierarchical Bayesian analysis of human dose response data. Wat. Res. 35, 3934–3940. Meuten, D.J., Van Kruiningen, H.J., and Lein, D.H. 1974. Cryptosporidiosis in a calf. J. Am. Vet. Med. Assoc. 165, 914–917. Millard, P.S., Gensheimer, K.F., Addiss, D.G., Sosin, D.M., Beckett, G.A., Houck-Jankoski, A., and Hudson, A. 1994. An outbreak of cryptosporidiosis from fresh-pressed apple cider. J. Am. Med. Assoc. 272, 1592–1596. Molbak, K., Hojlyng, N., Ingholt, L.A., da Silva, P., Jepsen, S., and Aaby, P. 1990. An epidemic outbreak of cryptosporidiosis, a prospective community study from Guinea Bissau. Pediatr. Infect. Dis. J. 9, 566–570. 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. Moodley, D., Jackson, T.F., Gathiram, V., and van den, Ende E.J. 1991. Cryptosporidium infections in children in Durban: Seasonal variation, age distribution and disease status. S. Afr. Med. J. 79, 295–297. Moore, A.C., Herwaldt, B.L., Craun, G.F., Calderon, R.L., Highsmith, A.K., and Juranek, D.D. 1993. Surveillance for waterborne disease outbreaks—United States, 1991–1992. MMWR CDC Surveill. Summ. 42, 1–22. Morgan, U., Weber, R., Xiao, L., Sulaiman, I., Thompson, R.C., Ndiritu, W., Lal, A., Moore, A., and Deplazes, P. 2000. 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-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. Moriarty, E.M., Duffy, G., McEvoy, J.M., Caccio, S., Sheridan, J.J., McDowell, D., and Blair, I.S. 2005. The effect of thermal treatments on the viability and infectivity of Cryptosporidium parvum on beef surfaces. J. Appl. Microbiol. 98, 618–623. Morin, M., Lariviere, S., and Lallier, R. 1976. Pathological and microbiological observations made on spontaneous cases of acute neonatal calf diarrhea. Can. J. Comp. Med. 40, 228–240. Morris, R.D., Naumova, E.N., and Griffiths, J.K. 1998. Did Milwaukee experience waterborne cryptosporidiosis before the large documented outbreak in 1993? Epidemiology 9, 264–270. Moss, D.M., Bennett, S.N., Arrowood, M.J., Wahlquist, S.P., and Lammie, P.J. 1998a. Enzyme-linked immunoelectrotransfer blot analysis of a cryptosporidiosis outbreak on a United States Coast Guard cutter. Am. J. Trop. Med. Hyg. 58, 110–118. Moss, D.M., Chappell, C.L., Okhuysen, P.C., DuPont, H.L., Arrowood, M.J., Hightower, A.W., and Lammie, P.J. 1998b. The antibody response to 27-, 17-, and 15-kDa Cryptosporidium antigens following experimental infection in humans. J. Infect. Dis. 178, 827–833. Moss, D.M., Montgomery, J.M., Newland, S.V., Priest, J.W., and Lammie, P.J. 2004. Detection of Cryptosporidium antibodies in sera and oral fluids using multiplex bead assay. J. Parasitol. 90, 397–404.


112


Mottet, C., El Wafa, A.A., Cavassini, M., D’Acremont, V., and Delarive, J. 2005. [Infectious diarrhea]. Rev. Med. Suisse. 1, 209–210, 213–217. 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. 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. Naumova, E.N., Christodouleas, J., Hunter, P.R., and Syed, Q. 2005. Effect of precipitation on seasonal variability in cryptosporidiosis recorded by the North West England surveillance system in 1990–1999. J. Wat. Health. 3, 185–196. Naumova, E.N., Egorov, A.I., Morris, R.D., and Griffiths, J.K. 2003. The elderly and waterborne Cryptosporidium infection: Gastroenteritis hospitalizations before and during the 1993 Milwaukee outbreak. Emerg. Infect. Dis. 9, 418–425. Navin, T.R. and Juranek, D.D. 1984. Cryptosporidiosis: Clinical, epidemiologic, and parasitologic review. Rev. Infect. Dis. 6, 313–327. Newman, R.D., Wuhib, T., Lima, A.A., Guerrant, R.L., and Sears, C.L. 1993. Environmental sources of Cryptosporidium in an urban slum in northeastern Brazil. Am. J. Trop. Med. Hyg. 49, 270–275. Nichols, G. 1992. The Biology, Epidemiology and Typing of Cryptosporidium spp. Ph.D. thesis. Surrey University. Nichols, G. 2003. Using existing surveillance based data. In Hunter P.R., Wait, M., and Ronchi, E., Eds., Drinking Water and Infectious Disease—Establishing the Links. CRC Press, Boca Raton, FL, pp. 131–141. Nichols, G. 2006. Infection risks from water in natural and man-made environments. Euro. Surveill. 11, 76–78. Nichols, G. and McLauchlin, J. 2003. Microbiology and the investigation of waterborne outbreaks: The use of Cryptosporidium typing in the investigation of waterborne disease. In Hunter, P.R., Wait, M., and Ronchi, E., Eds. Drinking Water and Infectious Disease—Establishing the Links. CRC Press, Boca Raton, FL, pp. 87–95. Nichols, G.L. 2000. Food-borne protozoa. Br. Med. Bull. 56, 209–235. Nichols, G.L. 2005. Fly transmission of Campylobacter. Emerg. Infect. Dis. 11, 361–364. Nichols, G.L., Chalmers, R.M., Sopwith, W., Regan, M., Hunter, C.A., Grenfell, P., Harrison, F., and Lane, C. 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. Nichols, G.L., McLauchlin, J., and Samuel, D. 1991. A technique for typing Cryptosporidium isolates. J. Protozool. 38, 237–240S. Nichols, R.A., Moore, J.E., and Smith, H.V. 2005. A rapid method for extracting oocyst DNA from Cryptosporidium-positive human feces for outbreak investigations. J. Microbiol. Meth. 65, 512–524. 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. Nydam, D.V. and Mohammed, H.O. 2005. Quantitative risk assessment of Cryptosporidium species infection in dairy calves. J. Dairy Sci. 88, 3932–3943. Okhuysen, P.C. 2001. Traveler’s diarrhea due to intestinal protozoa. Clin. Infect. Dis. 33, 110–114. Okhuysen, P.C. and Chappell, C.L. 2002. Cryptosporidium virulence determinants—are we there yet? Int. J. Parasitol. 32, 517–525. Okhuysen, P.C., Chappell, C.L., Crabb, J., Valdez, L.M., Douglass, E.T. and DuPont, H.L. 1998a. Prophylactic effect of bovine anti-Cryptosporidium hyperimmune colostrum immunoglobulin in healthy volunteers challenged with Cryptosporidium parvum. Clin. Infect. Dis. 26, 1324–1329. Okhuysen, P.C., Chappell, C.L., Crabb, J.H., Sterling, C.R., and DuPont, H.L. 1999. Virulence of three distinct Cryptosporidium parvum isolates for healthy adults. J. Infect. Dis. 180, 1275–1281. Okhuysen, P.C., Chappell, C.L., Sterling, C.R., Jakubowski, W., and DuPont, H.L. 1998b. Susceptibility and serologic response of healthy adults to reinfection with Cryptosporidium parvum. Infect. Immun. 66, 441–443. Okhuysen, P.C., Rich, S.M., Chappell, C.L., Grimes, K.A., Widmer, G., Feng, X., and Tzipori, S. 2002. Infectivity of a Cryptosporidium parvum isolate of cervine origin for healthy adults and interferon-gamma knockout mice. J. Infect. Dis. 185, 1320–1325.


Epidemiology

113

Okhuysen, P.C., Rogers, G.A., Crisanti, A., Spano, F., Huang, D.B., Chappell, C.L., and Tzipori, S. 2004. Antibody response of healthy adults to recombinant thrombospondin-related adhesive protein of Cryptosporidium 1 after experimental exposure to Cryptosporidium oocysts. Clin. Diagn. Lab. Immunol. 11, 235–238. Ong, C.S., Eisler, D.L., Alikhani, A., Fung, V.W., Tomblin, J., Bowie, W.R., and Isaac-Renton, J.L. 2002. Novel Cryptosporidium genotypes in sporadic cryptosporidiosis cases: First report of human infections with a cervine genotype. Emerg. Infect. Dis. 8, 263–268. 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. Palmer, C.J., Xiao, L., Terashima, A., Guerra, H., Gotuzzo, E., Saldias, 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. Park, J.H., Guk, S.M., Han, E.T., Shin, E.H., Kim, J.L., and Chai, J.Y. 2006. Genotype analysis of Cryptosporidium spp. prevalent in a rural village in Hwasun-gun, Republic of Korea. Kor. J. Parasitol. 44, 27–33. Patel, S., Pedraza-Diaz, S., and McLauchlin, J. 1999. The identification of Cryptosporidium species and Cryptosporidium parvum directly from whole feces by analysis of a multiplex PCR of the 18S rRNA gene and by PCR/RFLP of the Cryptosporidium outer wall protein (COWP) gene. Int. J. Parasitol. 29, 1241–1247. Patel, S., Pedraza-Diaz, S., McLauchlin, J., and Casemore, D.P. 1998. Molecular characterisation of Cryptosporidium parvum from two large suspected waterborne outbreaks. Outbreak Control Team South and West Devon 1995, Incident Management Team and Further Epidemiological and Microbiological Studies Subgroup North Thames 1997. Commun. Dis. Pub. Health. 1, 231–233. Pedersen, C., Danner, S., Lazzarin, A., Glauser, M.P., Weber, R., Katlama, C., Barton, S.E., and Lundgren, J.D. 1996. Epidemiology of cryptosporidiosis among European AIDS patients. Genitourin. Med. 72, 128–131. Pedraza-Diaz, S., Amar, C., Iversen, A.M., Stanley, P.J., and McLauchlin, J. 2001a. Unusual Cryptosporidium species recovered from human feces: First description of Cryptosporidium felis and Cryptosporidium “dog type” from patients in England. J. Med. Microbiol. 50, 293–296. Pedraza-Diaz, S., Amar, C., and McLauchlin, J. 2000. The identification and characterisation of an unusual genotype of Cryptosporidium from human feces as Cryptosporidium meleagridis. FEMS Microbiol. Lett. 189, 189–194. Pedraza-Diaz, S., Amar, C.F., 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 Youngs, E.R. 2001b. Cryptosporidium meleagridis from humans: Molecular analysis and description of affected patients. J. Infect. 42, 243–250. Peng, M.M., Meshnick, S.R., Cunliffe, N.A., Thindwa, B.D., Hart, C.A., Broadhead, R.L., and Xiao, L. 2003. Molecular epidemiology of cryptosporidiosis in children in Malawi. J. Eukaryot. Microbiol. 50 Suppl., 557–559. Peng, M.M., Xiao, L., Freeman, A.R., Arrowood, M.J., Escalante, A.A., Weltman, A.C., Ong, C.S., MacKenzie, W.R., Lal, A.A., and Beard, C.B. 1997. Genetic polymorphism among Cryptosporidium parvum isolates, evidence of two distinct human transmission cycles. Emerg. Infect. Dis. 3, 567–573. 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. Pieniazek, N.J., Bornay-Llinares, F.J., Slemenda, S.B., da Silva, A.J., Moura, I.N., Arrowood, M.J., Ditrich, O., and Addiss, D.G. 1999. New Cryptosporidium genotypes in HIV-infected persons. Emerg. Infect. Dis. 5, 444–449. 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. Pohjola, S., Oksanen, H., Jokipii, L., and Jokipii, A.M. 1986. Outbreak of cryptosporidiosis among veterinary students. Scand. J. Infect. Dis. 18, 173–178.


114


Pohlenz, J., Moon, H.W., Cheville, N.F., and Bemrick, W.J. 1978. Cryptosporidiosis as a probable factor in neonatal diarrhea of calves. J. Am. Vet. Med. Assoc. 172, 452–457. Preiser, G., Preiser, L., and Madeo, L. 2003. An outbreak of cryptosporidiosis among veterinary science students who work with calves. J. Am. Coll. Health. 51, 213–215. 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, 123–131. 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. Pritchard, G.C., Marshall, J.A., Giles, M., Chalmers, R., and Marshall, R.N. 2006. Cryptosporidium parvum infection in orphan lambs on a farm open to the public. Vet. Rec. Submitted. Pruss, A., Kay, D., Fewtrell, L., and Bartram, J. 2002. Estimating the burden of disease from water, sanitation, and hygiene at a global level. Environ. Health Perspect. 110, 537–542. Puech, M.C., McAnulty, J.M., Lesjak, M., Shaw, N., Heron, L., and Watson, J.M. 2001. A statewide outbreak of cryptosporidiosis in New South Wales associated with swimming at public pools. Epidemiol. Infect. 126, 389–396. Quiroz, E.S., Bern, C., MacArthur, J.R., Xiao, L., Fletcher, M., Arrowood, M.J., Shay, D.K., Levy, M.E., Glass, R.I., and A.Lal. 2000. An outbreak of cryptosporidiosis linked to a foodhandler. J. Infect. Dis. 181, 695–700. Rahman, A.S., Sanyal, S.C., Al Mahmud, K.A., and Sobhan, A. 1985. Cryptosporidium diarrhea in calves and their handlers in Bangladesh. Ind. J. Med. Res. 82, 510–516. Ravn, P., Lundgren, J.D., Kjaeldgaard, P., Holten-Anderson, W., Hojlyng, N., Nielsen, J.O., and Gaub, J. 1991. Nosocomial outbreak of cryptosporidiosis in AIDS patients. Br. Med. J. 302, 277–280. Reese, N.C., Current, W.L.. Ernst, J.V., and Bailey, W.S. 1982. Cryptosporidiosis of man and calf: A case report and results of experimental infections in mice and rats. Am. J. Trop. Med. Hyg. 31, 226–229. Reif, J.S., Wimmer, L., Smith, J.A., Dargatz, D.A., and Cheney. J.M. 1989. Human cryptosporidiosis associated with an epizootic in calves. Am. J. Pub. Health. 79, 1528–1530. Ribeiro, C.D. and Palmer, S.R. 1986. Family outbreak of cryptosporidiosis. Br. Med. J. (Clin. Res. Ed). 292, 377. Rice, G., Heberling, M.T., Rothermich, M., Wright, J.M., Murphy, P.A., Craun, M.F., and Craun, G.F. 2006. The role of disease burden measures in future estimates of endemic waterborne disease. J. Wat. Health. 4 (Suppl. 2), 187–199. Richardson, A.J., Frankenberg, R.A., Buck, A.C., Selkon, J.B., Colbourne, J.S., Parsons, J.W., and MayonWhite, R.T. 1991. An outbreak of waterborne cryptosporidiosis in Swindon and Oxfordshire. Epidemiol. Infect. 107, 485–495. Robertson, B., Sinclair, M.I., Forbes, A.B., Veitch, M., Kirk, M., Cunliffe, D., Willis, J., and Fairley, C.K. 2002. Case-control studies of sporadic cryptosporidiosis in Melbourne and Adelaide, Australia. Epidemiol. Infect. 128, 419–431. Robertson, L.J. and Gjerde, B. 2001a. Factors affecting recovery efficiency in isolation of Cryptosporidium oocysts and Giardia cysts from vegetables for standard method development. J. Food Prot. 64, 1799–1805. Robertson, L.J. and Gjerde, B. 2001b. Occurrence of parasites on fruits and vegetables in Norway. J. Food Prot. 64, 1793–1798. Robertson, L.J., Greig, J.D., Gjerde, B., and Fazil, A. 2005. The potential for acquiring cryptosporidiosis or giardiosis from consumption of mung bean sprouts in Norway, a preliminary step-wise risk assessment. Int. J. Food Microbiol. 98, 291–300. Robinson, P., Okhuysen, P.C., Chappell, C.L., Lewis, D.E., Shahab, I., Janecki, A., and White, Jr., A.C. 2001a. 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., Lewis, D.E., Shahab, I., Lahoti, S., and White, Jr., A.C. 2000. Transforming growth factor beta1 is expressed in the jejunum after experimental Cryptosporidium parvum infection in humans. Infect. Immun. 68, 5405–5407.


Epidemiology

115

Robinson, P., Okhuysen, P.C., Chappell, C.L., Lewis, D.E., Shahab, I., Lahoti, S., and White, Jr., A.C. 2001b. Expression of IL-15 and IL-4 in IFN-gamma-independent control of experimental human Cryptosporidium parvum infection. Cytokine 15, 39–46. Robinson, P., Okhuysen, P.C., Chappell, C.L., Weinstock, J.V., Lewis, D.E., Actor, J.K., and White, Jr., A.C. 2003. Substance P expression correlates with severity of diarrhea in cryptosporidiosis. J. Infect. Dis. 188, 290–296. Rodrigues, F., Davies, E.G., Harrison, P., McLauchlin, J., Karani, J., Portmann, B., Jones, A., Veys, P., MieliVergani, G., and Hadzic, N. 2004. Liver disease in children with primary immunodeficiencies. J. Pediatr. 145, 333–339. 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. Rodriguez-Salinas, P.E., Aragon Pena, A.J., Allue, T.M., Lopaz Perez, M.A., Jimenez, M.M., and Dominguez Rodriguez, M.J. 2000. [Outbreak of cryptosporidiosis in Guadarrama (Autonomous Community of Madrid)]. Rev. Esp. Salud Publica 74, 527–536. Rooney, R.M., Bartram, J.K., Cramer, E.H., Mantha, S., Nichols, G., Suraj, R., and Todd, E.C. 2004a. A review of outbreaks of waterborne disease associated with ships: Evidence for risk management. Pub. Health. Rep. 119, 435–442. Rooney, R.M., Cramer, E.H., Mantha, S., Nichols, G., Bartram, J.K., Farber, J.M., and Benembarek, P.K. 2004b. A review of outbreaks of foodborne disease associated with passenger ships: Evidence for risk management. Pub. Health. Rep. 119, 427–434. Rose, J.B. and Slifko, T.R. 1999. Giardia, Cryptosporidium, and Cyclospora and their impact on foods: A review. J. Food Prot.. 62, 1059–1070. Roy, S.L., DeLong, S.M., Stenzel, S.A., Shiferaw, B., Roberts, J.M., Khalakdina, A., Marcus, R., Segler, S.D., Shah, D.D., Thomas, S., Vugia, D.J., Zansky, S.M., Dietz, V., and Beach, M.J. 2004. Risk factors for sporadic cryptosporidiosis among immunocompetent persons in the United States from 1999 to 2001. J. Clin. Microbiol. 42, 2944–2951. Roy, S.L., Scallan, E., and Beach, M.J. 2006. The rate of acute gastrointestinal illness in developed countries. J. Wat. Health. 4 (Suppl. 2), 31–69. Rush, B.A., Chapman, P.A., and Ineson, R.W. 1990. A probable waterborne outbreak of cryptosporidiosis in the Sheffield area. J. Med. Microbiol. 32, 239–242. Sacco, C., Bianchi, M., Lorini, C., Burrini, D., Berchielli, S., and Lanciotti, E. 2006. [Removal of Cryptosporidium and Giardia in drinking water treatment in a Tuscan area]. Ann. Ig. 18, 117–126. Said, B., Wright F., Nichols, G.L., Reacher, M., and Rutter, M. 2003. Outbreaks of infectious disease associated with private drinking water supplies in England and Wales 1970–2000. Epidemiol. Infect. 130, 469–479. Sayers, G.M., Dillon, M.C., Connolly, E., Thornton, L., Hyland, E., Loughman, E., O’Mahony, M.A., and Butler, K.M. 1996. Cryptosporidiosis in children who visited an open farm. Commun. Dis. Rep. CDR Rev. 6, R140–R144. Semenza, J. and Nichols, G. 2007. Cryptosporidiosis surveillance and water-borne outbreaks in Europe. Eurosurveillance, 12 (5), 1–8. Slapeta, J. 2006. Cryptosporidium species found in cattle: A proposal for a new species. Trends Parasitol. 22, 469–474. Slapeta, J. 2007. Response to Xiao et al., Further debate on the description of Cryptosporidium pestis. Trends Parasitol. 23, 42–43. Smerdon, W.J., Nichols, T., Chalmers, R.M., Heine, H., and Reacher, M.H. 2003. Foot and mouth disease in livestock and reduced cryptosporidiosis in humans, England and Wales. Emerg. Infect. Dis. 9, 22–28. Smith, A., Reacher, M., Smerdon, W., Adak, G.K., Nichols, G., and Chalmers, R.M. 2006. Outbreaks of waterborne infectious intestinal disease in England and Wales, 1992–2003. Epidemiol. Infect. 134, 1141–1149. Smith, H.V., Girdwood, R.W., Patterson, W.J., Hardie, R., Green, L.A., Benton, C., Tulloch, W., Sharp, J.C., and Forbes, G.I. 1988. Waterborne outbreak of cryptosporidiosis. Lancet 2, 1484. Smith, H.V., Nichols, R.A., Mallon, M., Macleod, A., Tait A., Reilly, W.J., Browning, L.M., Gray, D., Reid, S.W., and Wastling, J.M. 2005. Natural Cryptosporidium hominis infections in Scottish cattle. Vet. Rec. 156, 710 and 711.


116


Smith, H.V., Patterson, W.J., Hardie, R., Greene, L.A., Benton, C., Tulloch, W., Gilmour, R.A., Girdwood, R.W., Sharp, J.C., and Forbes, G.I. 1989. An outbreak of waterborne cryptosporidiosis caused by posttreatment contamination. Epidemiol. Infect. 103, 703–715. Smith, K.E., Stenzel, S.A., Bender J.B., Wagstrom, E., Soderlund, D., Leano, F.T., Taylor, C.M., Belle-Isle, P.A., and Danila, R. 2004. Outbreaks of enteric infections caused by multiple pathogens associated with calves at a farm day camp. Pediatr. Infect. Dis. J. 23, 1098–1104. Soave, R. and Armstrong, D. 1986. Cryptosporidium and cryptosporidiosis. Rev. Infect. Dis. 8, 1012–1023. 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. Soller, J.A. 2006. Use of microbial risk assessment to inform the national estimate of acute gastrointestinal illness attributable to microbes in drinking water. J. Wat. Health. 4 (Suppl. 2), 165–186. 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. Spano, F., Putignani, L., Crisanti, A., Sallicandro, P., Morgan, U.M., Le Blancq, S.M., Tchack, L., Tzipori, S., and Widmer, G. 1998a. Multilocus genotypic analysis of Cryptosporidium parvum isolates from different hosts and geographical origins. J. Clin. Microbiol. 36, 3255–3259. Spano, F., Putignani, L., Guida, S., and Crisanti, A. 1998b. Cryptosporidium parvum, PCR-RFLP analysis of the TRAP-C1 (thrombospondin-related adhesive protein of Cryptosporidium-1) gene discriminates between two alleles differentially associated with parasite isolates of animal and human origin. Exp. Parasitol. 90, 195–198. Spano, F., Putignani, L., McLauchlin, J., Casemore, D.P., and Crisanti, A. 1997. PCR-RFLP analysis of the Cryptosporidium oocyst wall protein (COWP) gene discriminates between C. wrairi and C. parvum, and between C. parvum isolates of human and animal origin. FEMS Microbiol. Lett. 150, 209–217. Squier, C., Yu, V.L., and Stout, J.E. 2000. Waterborne Nosocomial Infections. Curr Infect. Dis. Rep. 2, 490–496. 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. Stantic-Pavlinic, M., Xiao, L., Glaberman, S., Lal, A.A., Orazen, T., Rataj-Verglez, A., Logar, J., and Berce, I. 2003b. Cryptosporidiosis associated with animal contacts. Wien. Klin. Wochenschr. 115, 125–127. Stantic-Pavlinic, M., Xiao, L., Glaberman, S., Lal, A.A., Orazen, T., Rataj-Verglez, A., Logar, J., and Berce, I. 2003a. Cryptosporidiosis associated with animal contacts. Wien. Klin. Wochenschr. 115, 125–127. Steele, A.D., Gove, E., and Meewes, P.J. 1989. Cryptosporidiosis in white patients in South Africa. J. Infect. 19, 281–285. Stefanogiannis, N., McLean, M., and Van Mil, H. 2001. Outbreak of cryptosporidiosis linked with a farm event. N. Z. Med. J. 114, 519–521. Steffen, R., Mathewson, J.J., Ericsson, C.D., DuPont, H.L., Helminger, A., Balm, T.K., Wolff, K., and Witassek, F. 1988. Travelers’ diarrhea in West Africa and Mexico: Fecal transport systems and liquid bismuth subsalicylate for self-therapy. J. Infect. Dis. 157, 1008–1013. Sterling, C.R. and Arrowood, M.J. 1986. Detection of Cryptosporidium sp. infections using a direct immunofluorescent assay. Pediatr. Infect. Dis. 5, S139–S142. Sterling, C.R., Seegar, K., and Sinclair, N.A. 1986. Cryptosporidium as a causative agent of traveler’s diarrhea. J. Infect. Dis. 153, 380 and 381. Stirling, R., Aramini, J., Ellis, A., Lim, G., Meyers, R., Fleury, M., and Werker, D. 2001. Waterborne cryptosporidiosis outbreak, North Battleford, Saskatchewan, Spring 2001. Can. Commun. Dis. Rep. 27, 185–192. Strachan, N.J., Ogden, I.D., Smith-Palmer, A., and Jones, K. 2003. Foot and mouth epidemic reduces cases of human cryptosporidiosis in Scotland. J. Infect. Dis. 188, 783–786. Strohmeyer, R.A., Morley, P.S., Hyatt, D.R., Dargatz, D.A., Scorza, A.V., and Lappin, M.R. 2006. Evaluation of bacterial and protozoal contamination of commercially available raw meat diets for dogs. J. Am. Vet. Med. Assoc. 228, 537–542.


Epidemiology

117

Sulaiman, I.M., Morgan, U.M., Thompson, R.C., Lal, A.A., and Xiao, L. 2000. Phylogenetic relationships of Cryptosporidium parasites based on the 70-kilodalton heat shock protein (HSP70) gene. Appl. Environ. Microbiol. 66, 2385–2391. 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, R190–R192. Sutthikornchai, C., Jantanavivat, C., Thongrungkiat, S., Harnroongroj, T., and Sukthana, Y. 2005. Protozoal contamination of water used in Thai frozen food industry. SE Asian J. Trop. Med. Pub. Health. 36 (Suppl. 4), 41–45. Szostakowska, B., Kruminis-Lozowska, W., Racewicz, M., Knight, R., Tamang, L., Myjak, P., and Graczyk, T.K. 2004. Cryptosporidium parvum and Giardia lamblia recovered from flies on a cattle farm and in a landfill. Appl. Environ. Microbiol. 70, 3742–3744. 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. Tanriverdi, S. and Tzipori, S. 2006. Cryptosporidium hominis, experimental challenge of healthy adults. Am. J. Trop. Med. Hyg. 75, 851–857. Tanriverdi, S. and Widmer, G. 2006. Differential evolution of repetitive sequences in Cryptosporidium parvum and Cryptosporidium hominis. Infect. Genet. Evol. 6, 113–122. Teresa, O.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. Teunis, P.F., Chappell, C.L., and Okhuysen, P.C. 2002a. Cryptosporidium dose response studies: Variation between isolates. Risk Anal. 22, 175–183. Teunis, P.F., Chappell, C.L., and Okhuysen, P.C. 2002b. Cryptosporidium dose-response studies, Variation between hosts. Risk Anal. 22, 475–485. Thomas, K.M., Charron, D.F., Waltner-Toews, D., Schuster, C., Maarouf, A.R., and Holt, J.D. 2006. A role of high impact weather events in waterborne disease outbreaks in Canada, 1. Int. J. Environ. Health. Res. 16, 167–180. Thurston-Enriquez, J.A., Watt, P., Dowd, S.E., Enriquez, R., Pepper, I.L., and Gerba. C.P. 2002. Detection of protozoan parasites and microsporidia in irrigation waters used for crop production. J. Food Prot. 65, 378–382. Tiangtip, R. and Jongwutiwes, S. 2002. Molecular analysis of Cryptosporidium species isolated from HIVinfected patients in Thailand. Trop. Med. Int. Health. 7, 357–364. 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. Trotz-Williams, L.A., Martin, D.S., Gatei, W., Cama, V., Peregrine, A.S., Martin, S.W., Nydam, D.V., Jamieson, F., and Xiao, L. 2006. Genotype and subtype analyses of Cryptosporidium isolates from dairy calves and humans in Ontario. Parasitol. Res. 99, 346–352. 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. Tzipori, S. and Campbell, I. 1981. Prevalence of Cryptosporidium antibodies in 10 animal species. J. Clin. Microbiol. 14, 455 and 456. Ungar, B.L., Mulligan, M., and Nutman, T.B. 1989. Serologic evidence of Cryptosporidium infection in US volunteers before and during Peace Corps service in Africa. Arch. Intern. Med. 149, 894–897. Ungar, B.L. and Nash, T.E. 1986. Quantification of specific antibody response to Cryptosporidium antigens by laser densitometry. Infect. Immun. 53, 124–128. Ungar, B.L., Soave, R., Fayer, R., and Nash, T.E. 1986. Enzyme immunoassay detection of immunoglobulin M and G antibodies to Cryptosporidium in immunocompetent and immunocompromised persons. J. Infect. Dis. 153, 570–578. Westrell, T., Schonning, C., Stenstrom, T.A., and Ashbolt, N.J. 2004. QMRA (quantitative microbial risk assessment) and HACCP (hazard analysis and critical control points) for management of pathogens in wastewater and sewage sludge treatment and reuse. Wat. Sci. Technol. 50, 23–30. White, A.C., Robinson, P., Okhuysen, P.C., Lewis, D.E., Shahab, I., Lahoti, S., DuPont, H.L., and Chappell, C.L. 2000. Interferon-gamma expression in jejunal biopsies in experimental human cryptosporidiosis correlates with prior sensitization and control of oocyst excretion. J. Infect. Dis. 181, 701–709. Wilberschied, L. 1995. A swimming-pool-associated outbreak of cryptosporidiosis. Kans. Med. 96, 67 and 68.


118


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.


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.

119


120


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


Molecular Epidemiology

121

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).


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



123

(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,


124

Cryptosporidium and Cryptosporidiosis, Second Edition 0.1 substitutions/site 100

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



125

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


126


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.



127

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


128


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



129

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


130


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


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

Intestinal

0.02 substitutions/site

131

Gastric


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).


132


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.



133

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


134


(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


Molecular Epidemiology 135

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



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




138


(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).



139

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


140


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).



141

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

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


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



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




144


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.



145

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).


146


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



147

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



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




150


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.



151

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.

VIII. References Abe, N., Matsubayashi, M., Kimata, I., and Iseki, M. 2006. Subgenotype analysis of Cryptosporidium parvum isolates from humans and animals in Japan using the 60-kDa glycoprotein gene sequences. Parasitol. Res. 99, 303–305. Abrahamsen, M.S., Templeton, T.J., Enomoto, S., Abrahante, J.E., Zhu, G., Lancto, C.A., Deng, M., Liu, C., Widmer, G., Tzipori, S., Buck, G.A., Xu, P., Bankier, A.T., Dear, P.H., Konfortov, B.A., Spriggs, H.F., Iyer, L., Anantharaman, V., Aravind, L., and Kapur, V. 2004. Complete genome sequence of the apicomplexan, Cryptosporidium parvum. Science. 304, 441–445. Aiello, A.E., Xiao, L., Limor, J.R., Liu, C., Abrahamsen, M.S., and Lal, A.A. 1999. Microsatellite analysis of the human and bovine genotypes of Cryptosporidium parvum. J. Eukaryot. Microbiol. 46, 46S and 47S. 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, 1828–32. Akiyoshi, D.E., Tumwine, J.K., Bakeera-Kitaka, S., and Tzipori, S. 2006. Subtype analysis of Cryptosporidium isolates from children in Uganda. J. Parasitol. 92, 1097–1100. Almeida, A.A., Delgado, M.L., Soares, S.C., Castro, A.O., Moreira, M.J., Mendonca, C.M., Canada, N.B., Correia Da Costa, J.M., and Coelho, H.G. 2006. Genetic characterization of Cryptosporidium isolates from humans in Northern Portugal. J. Eukaryot. Microbiol. 53, S26 and S27. Alves, M., Matos, O., and Antunes, F. 2001. Multilocus PCR-RFLP analysis of Cryptosporidium isolates from HIV-infected patients from Portugal. Ann. Trop. Med. Parasitol. 95, 627–632. Alves, M., Matos, O., and Antunes, F. 2003a. Microsatellite analysis of Cryptosporidium hominis and C. parvum in Portugal: A preliminary study. J. Eukaryot. Microbiol. 50 Suppl., 529 and 530. Alves, M., Ribeiro, A.M., Neto, C., Ferreira, E., Benoliel, M.J., Antunes, F., and Matos, O. 2006a. Distribution of Cryptosporidium species and subtypes in water samples in Portugal: A preliminary study. J. Eukaryot. Microbiol. 53, S24 and S25. Alves, M., Xiao, L., Antunes, F., and Matos, O. 2006b. Distribution of Cryptosporidium subtypes in humans and domestic and wild ruminants in Portugal. Parasitol. Res. 99, 287–292. Alves, M., Xiao, L., Sulaiman, I., Lal, A.A., Matos, O., and Antunes, F. 2003b. Subgenotype analysis of Cryptosporidium isolates from humans, cattle, and zoo ruminants in Portugal. J. Clin. Microbiol. 41, 2744–2747. Asahi, H., Koyama, T., Arai, H., Funakoshi, Y., Yamaura, H., Shirasaka, R., and Okutomi, K. 1991. Biological nature of Cryptosporidium sp. isolated from a cat. Parasitol. Res. 77, 237–240. Atwill, E.R., Phillips, R., Pereira, M.D., Li, X., and McCowan, B. 2004. Seasonal shedding of multiple Cryptosporidium genotypes in California ground squirrels (Spermophilus beecheyi). Appl. Environ. Microbiol. 70, 6748–6752.


152


Awad-el-Kariem, F.M., Warhurst, D.C., and McDonald, V. 1994. Detection and species identification of Cryptosporidium oocysts using a system based on PCR and endonuclease restriction. Parasitology 109, 19–22. Baishanbo, A., Gargala, G., Delaunay, A., Francois, A., Ballet, J.J., and Favennec, L. 2005. Infectivity of Cryptosporidium hominis and Cryptosporidium parvum genotype 2 isolates in immunosuppressed Mongolian gerbils. Infect. Immun. 73, 5252–5255. Bajer, A., Caccio, S., Bednarska, M., Behnke, J.M., Pieniazek, N.J., and Sinski, E. 2003. Preliminary molecular characterization of Cryptosporidium parvum isolates of wildlife rodents from Poland. J. Parasitol. 89, 1053–1055. Bauer, D. 1994. The capacity of dogs to serve as reservoirs for gastrointestinal disease in children. Ir. Med. J. 87, 184 and 185. Bednarska, M., Bajer, A., Kulis, K., and Sinski, E. 2003. Biological characterization of Cryptosporidium parvum isolates of wildlife rodents in Poland. Ann. Agric. Environ. Med. 10, 163–169. Bjorkman, C. and Mattsson, J.G. 2006. Persistent infection in a dairy herd with an unusual genotype of bovine Cryptosporidium parvum. FEMS Microbiol. Lett. 254, 71–74. Blackburn, B.G., Mazurek, J.M., Hlavsa, M., Park, J., Tillapaw, M., Parrish, M., Salehi, E., Franks, W., Koch, E., Smith, F., Xiao, L., Arrowood, M., Hill, V., da Silva, A., Johnston, S., and Jones, J.L. 2006. Cryptosporidiosis associated with ozonated apple cider. Emerg. Infect. Dis. 12, 684–686. Bonnin, A., Fourmaux, M.N., Dubremetz, J.F., Nelson, R.G., Gobet, P., Harly, G., Buisson, M., PuygauthierToubas, D., Gabriel-Pospisil, G., Naciri, M., and Camerlynck, P. 1996. Genotyping human and bovine isolates of Cryptosporidium parvum by polymerase chain reaction-restriction fragment length polymorphism analysis of a repetitive DNA sequence. FEMS Microbiol. Lett. 137, 207–211. Bushen, O.Y., Kohli, A., Pinkerton, R.C., Dupnik, K., Newman, R.D., Sears, C.L., Fayer, R., Lima, A.A., and Guerrant, R.L. 2007. Heavy cryptosporidial infections in children in northeast Brazil: Comparison of Cryptosporidium hominis and Cryptosporidium parvum. Trans. R. Soc. Trop. Med. Hyg. 101, 378–384. Caccio, S., Homan, W., Camilli, R., Traldi, G., Kortbeek, T., and Pozio, E. 2000. A microsatellite marker reveals population heterogeneity within human and animal genotypes of Cryptosporidium parvum. Parasitology 120, 237–244. Caccio, S., Homan, W., van Dijk, K., and Pozio, E. 1999. Genetic polymorphism at the beta-tubulin locus among human and animal isolates of Cryptosporidium parvum. FEMS Microbiol. Lett. 170, 173–179. Caccio, S., Pinter, E., Fantini, R., Mezzaroma, I., and Pozio, E. 2002. Human infection with Cryptosporidium felis: Case report and literature review. Emerg. Infect. Dis. 8, 85 and 86. Caccio, S., Spano, F., and Pozio, E. 2001. Large sequence variation at two microsatellite loci among zoonotic (genotype C) isolates of Cryptosporidium parvum. Int. J. Parasitol. 31, 1082–1086. Caccio, S.M. 2005. Molecular epidemiology of human cryptosporidiosis. Parasitologia. 47, 185–192. Cama, V., Gilman, R.H., Vivar, A., Ticona, E., Ortega, Y., Bern, C., and Xiao, L. 2006a. Mixed Cryptosporidium infections and HIV. Emerg. Infect. Dis. 12, 1025–1028. Cama, V.A., Arrowood, M.J., Ortega, Y.R., and Xiao, L. 2006b. Molecular characterization of the Cryptosporidium parvum IOWA isolate kept in different laboratories. J. Eukaryot. Microbiol. 53, S40–S42. Cama, V.A., Bern, C., Sulaiman, I.M., Gilman, R.H., Ticona, E., Vivar, A., Kawai, V., Vargas, D., Zhou, L., and Xiao, L. 2003. Cryptosporidium species and genotypes in HIV-positive patients in Lima, Peru. J. Eukaryot. Microbiol. 50 Suppl., 531–533. Cama, V.A., Ross, J.M., Crawford, S., Kawai, V., Chavez-Valdez, R., Vargas, D., Vivar, A., Ticona, E., Ñavincopa, M., Williamson, J., Ortega, Y., Gilman, R.H., Bern, C., and Xiao, L. 2007. Differences in clinical manifestations among Cryptosporidium species and subtypes in HIV-infected persons. J. Infect. Dis. (in press). Certad, G., Ngouanesavanh, T., Hernan, A., Rojas, E., Contreras, R., Pocaterra, L., Nunez, L., Dei-Cas, E., and Guyot, K. 2006. First molecular data on cryptosporidiosis in Venezuela. J. Eukaryot. Microbiol. 53, S30–S32. Cevallos, A.M., Zhang, X., Waldor, M.K., Jaison, S., Zhou, X., Tzipori, S., Neutra, M.R., and Ward, H.D. 2000. Molecular cloning and expression of a gene encoding Cryptosporidium parvum glycoproteins gp40 and gp15. Infect. Immun. 68, 4108–4116. Chalmers, R.M., Elwin, K., Thomas, A.L., and Joynson, D.H. 2002. Infection with unusual types of Cryptosporidium is not restricted to immunocompromised patients. J. Infect. Dis. 185, 270 and 271.



153

Chalmers, R.M., Ferguson, C., Caccio, S., Gasser, R.B., Abs El-Osta, Y.G., Heijnen, L., Xiao, L., Elwin, K., Hadfield, S., Sinclair, M., and Stevens, M. 2005a. Direct comparison of selected methods for genetic categorization of Cryptosporidium parvum and Cryptosporidium hominis species. Int. J. Parasitol. 35, 397–410. Chalmers, R.M., Thomas, A.L., Butler, B.A., and Morel, M.C. 2005b. Identification of Cryptosporidium parvum genotype 2 in domestic horses. Vet. Rec. 156, 49 and 50. Chappell, C.L., Okhuysen, P.C., Langer-Curry, R., Widmer, G., Akiyoshi, D.E., Tanriverdi, S., and Tzipori, S. 2006. Cryptosporidium hominis: Experimental challenge of healthy adults. Am. J. Trop. Med. Hyg. 75, 851–857. Chung, P.A., Johnson, J., Khramtsov, N.V., and Upton, S.J. 2000. Cloning and molecular characterization of a gene encoding a Cryptosporidium parvum putative 20S proteasome beta1-type subunit. DNA Seq. 11, 309–314. Cohen, S., Dalle, F., Gallay, A., Di Palma, M., Bonnin, A., and Ward, H.D. 2006. Identification of Cpgp40/15 type Ib as the predominant allele in isolates of Cryptosporidium spp. from a waterborne outbreak of gastroenteritis in South Burgundy, France. J. Clin. Microbiol. 44, 589–591. Coupe, S., Delabre, K., Pouillot, R., Houdart, S., Santillana-Hayat, M., and Derouin, F. 2006. Detection of Cryptosporidium, Giardia and Enterocytozoon bieneusi in surface water, including recreational areas: A one-year prospective study. FEMS Immunol. Med. Microbiol. 47, 351–359. Coupe, S., Sarfati, C., Hamane, S., and Derouin, F. 2005. Detection of Cryptosporidium and identification to the species level by nested PCR and restriction fragment length polymorphism. J. Clin. Microbiol. 43, 1017–1023. da Silva, A.J., Caccio, S., Williams, C., Won, K.Y., Nace, E.K., Whittier, C., Pieniazek, N.J., and Eberhard, M.L. 2003. Molecular and morphologic characterization of a Cryptosporidium genotype identified in lemurs. Vet. Parasitol. 111, 297–307. Das, P., Roy, S.S., Mitradhar, K., Dutta, P., Bhattacharya, M.K., Sen, A., Ganguly, S., Bhattacharya, S.K., Lal, A.A., and Xiao, L. 2006. Molecular characterization of Cryptosporidium spp. from children in Kolkata, India. J. Clin. Microbiol. 44, 4246–4249. Di Giovanni, G.D., Hashemi, F.H., Shaw, N.J., Abrams, F.A., LeChevallier, M.W., and Abbaszadegan, M. 1999. Detection of infectious Cryptosporidium parvum oocysts in surface and filter backwash water samples by immunomagnetic separation and integrated cell culture-PCR. Appl. Environ. Microbiol. 65, 3427–3432. Duke, L.A., Breathnach, A.S., Jenkins, D.R., Harkis, B.A., and Codd, A.W. 1996. A mixed outbreak of Cryptosporidium and campylobacter infection associated with a private water supply. Epidemiol. Infect. 116, 303–308. Egyed, Z., Sreter, T., Szell, Z., and Varga, I. 2003. Characterization of Cryptosporidium spp.—recent developments and future needs. Vet. Parasitol. 111, 103–114. Enemark, H.L., Ahrens, P., Juel, C.D., Petersen, E., Petersen, R.F., Andersen, J.S., Lind, P., and Thamsborg, S.M. 2002. Molecular characterization of Danish Cryptosporidium parvum isolates. Parasitology 125, 331–341. Enriquez, C., Nwachuku, N., and Gerba, C.P. 2001. Direct exposure to animal enteric pathogens. Rev. Environ. Health. 16, 117–131. Fayer, R., Santín, M., Trout, J.M., and Greiner, E. 2006. Prevalence of species and genotypes of Cryptosporidium found in 1-2-year-old dairy cattle in the eastern United States. Vet. Parasitol. 135, 105–112. Fayer, R., Santín, M., and Xiao, L. 2005. Cryptosporidium bovis n. sp. (Apicomplexa: Cryptosporidiidae) in cattle (Bos taurus). J. Parasitol. 91, 624–629. Feltus, D.C., Giddings, C.W., Schneck, B.L., Monson, T., Warshauer, D., and McEvoy, J.M. 2006. Evidence supporting zoonotic transmission of Cryptosporidium in Wisconsin. J. Clin. Microbiol. 44, 4303–4308. Feng, X., Rich, S.M., Akiyoshi, D., Tumwine, J.K., Kekitiinwa, A., Nabukeera, N., Tzipori, S., and Widmer, G. 2000. Extensive polymorphism in Cryptosporidium parvum identified by multilocus microsatellite analysis. Appl. Environ. Microbiol. 66, 3344–3349. Feng, X., Rich, S.M., Tzipori, S., and Widmer, G. 2002. Experimental evidence for genetic recombination in the opportunistic pathogen Cryptosporidium parvum. Mol. Biochem. Parasitol. 119, 55–62. Feng, Y., Ortega, Y., He, G., Das, P., Xu, M., Zhang, X., Fayer, R., Gatei, W., Cama, V., and Xiao, L. 2007. Wide geographic distribution of Cryptosporidium bovis and the deer-like genotype in bovines. Vet. Parasitol. (in press).


154


Fretz, R., Svoboda, P., Ryan, U.M., Thompson, R.C., Tanners, M., and Baumgartner, A. 2003. Genotyping of Cryptosporidium spp. isolated from human stool samples in Switzerland. Epidemiol. Infect. 131, 663–667. Gasser, R.B., Abs, E.L.O.Y.G., Prepens, S., and Chalmers, R.M. 2004. An improved “cold SSCP” method for the genotypic and subgenotypic characterization of Cryptosporidium. Mol. Cell Probes. 18, 329–332. Gasser, R.B., El-Osta, Y.G., and Chalmers, R.M. 2003. Electrophoretic analysis of genetic variability within Cryptosporidium parvum from imported and autochthonous cases of human cryptosporidiosis in the United Kingdom. Appl. Environ. Microbiol. 69, 2719–2730. Gatei, W., Ashford, R.W., Beeching, N.J., Kamwati, S.K., Greensill, J., and Hart, C.A. 2002a. Cryptosporidium muris infection in an HIV-infected adult, Kenya. Emerg. Infect. Dis. 8, 204–206. Gatei, W., Das, P., Dutta, P., Sen, A., Cama, V., Lal, A.A., and Xiao, L. 2007. Multilocus sequence typing and genetic structure of Cryptosporidium hominis from children in Kolkata, India. Infect. Genet. Evol. 7, 197–205. Gatei, W., Greensill, J., Ashford, R.W., Cuevas, L.E., Parry, C.M., Cunliffe, N.A., Beeching, N.J., and Hart, C.A. 2003. Molecular analysis of the 18S rRNA gene of Cryptosporidium parasites from patients with or without human immunodeficiency virus infections living in Kenya, Malawi, Brazil, the United Kingdom, and Vietnam. J. Clin. Microbiol. 41, 1458–1462. Gatei, W., Hart, C.A., Gilman, R.H., Das, P., Cama, V., and Xiao, L. 2006a. Development of a multilocus sequence typing tool for Cryptosporidium hominis. J. Eukaryot. Microbiol. 53, S43–S48. Gatei, W., Suputtamongkol, Y., Waywa, D., Ashford, R.W., Bailey, J.W., Greensill, J., Beeching, N.J., and Hart, C.A. 2002b. Zoonotic species of Cryptosporidium are as prevalent as the anthroponotic in HIVinfected patients in Thailand. Ann. Trop. Med. Parasitol. 96, 797–802. Gatei, W., Wamae, C.N., Mbae, C., Waruru, A., Mulinge, E., Waithera, T., Gatika, S.M., Kamwati, S.K., Revathi, G., and Hart, C.A. 2006b. Cryptosporidiosis: prevalence, genotype analysis, and symptoms associated with infections in children in Kenya. Am. J. Trop. Med. Hyg. 75, 78–82. Gibbons-Matthews, C. and Prescott, A.M. 2003. Intra-isolate variation of Cryptosporidium parvum small subunit ribosomal RNA genes from human hosts in England. Parasitol. Res. 90, 439–444. Gibbons, C.L., Ong, C.S., Miao, Y., Casemore, D.P., Gazzard, B.G., and Awad-El-Kariem, F.M. 2001. PCRELISA: A new simplified tool for tracing the source of cryptosporidiosis in HIV-positive patients. Parasitol. Res. 87, 1031–1034. Giles, M., Webster, K.A., Marshall, J.A., Catchpole, J., and Goddard, T.M. 2001. Experimental infection of a lamb with Cryptosporidium parvum genotype 1. Vet. Rec. 149, 523–525. Glaberman, S., Moore, J.E., Lowery, C.J., Chalmers, R.M., Sulaiman, I., Elwin, K., Rooney, P.J., Millar, B.C., Dooley, J.S., Lal, A.A., and Xiao, L. 2002. Three drinking-water-associated cryptosporidiosis outbreaks, Northern Ireland. Emerg. Infect. Dis. 8, 631–633. Glaberman, S., Sulaiman, I.M., Bern, C., Limor, J., Peng, M.M., Morgan, U., Gilman, R., Lal, A.A., and Xiao, L. 2001. A multilocus genotypic analysis of Cryptosporidium meleagridis. J. Eukaryot. Microbiol. 48 Suppl., 19S–22S. Glaeser, C., Grimm, F., Mathis, A., Weber, R., Nadal, D., and Deplazes, P. 2004. Detection and molecular characterization of Cryptosporidium spp. isolated from diarrheic children in Switzerland. Pediatr. Infect. Dis. J. 23, 359–361. Glaser, C.A., Safrin, S., Reingold, A., and Newman, T.B. 1998. Association between Cryptosporidium infection and animal exposure in HIV-infected individuals. J. Acquir. Immune Defic. Syndr. Hum. Retrovirol. 17, 79–82. Goh, S., Reacher, M., Casemore, D.P., Verlander, N.Q., Chalmers, R., Knowles, M., Williams, J., Osborn, K., and Richards, S. 2004. Sporadic cryptosporidiosis, North Cumbria, England, 1996–2000. Emerg. Infect. Dis. 10, 1007–1015. Goncalves, E.M., da Silva, A.J., Eduardo, M.B., Uemura, I.H., Moura, I.N., Castilho, V.L., and Corbett, C.E. 2006. Multilocus genotyping of Cryptosporidium hominis associated with diarrhea outbreak in a day care unit in Sao Paulo. Clinics 61, 119–126. Graczyk, T.K., Fayer, R., Trout, J.M., Lewis, E.J., Farley, C.A., Sulaiman, I., and Lal, A.A. 1998. Giardia sp. cysts and infectious Cryptosporidium parvum oocysts in the feces of migratory Canada geese (Branta canadensis). Appl. Environ. Microbiol. 64, 2736–2738. Grinberg, A., Oliver, L., Learmonth, J.J., Leyland, M., Roe, W., and Pomroy, W.E. 2003. Identification of Cryptosporidium parvum “cattle” genotype from a severe outbreak of neonatal foal diarrhea. Vet. Rec. 153, 628–631.



155

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. Guyot, K., Follet-Dumoulin, A., Lelievre, E., Sarfati, C., Rabodonirina, M., Nevez, G., Cailliez, J.C., Camus, D., and Dei-Cas, E. 2001. Molecular characterization of Cryptosporidium isolates obtained from humans in France. J. Clin. Microbiol. 39, 3472–3480. Hajdusek, O., Ditrich, O., and Slapeta, J. 2004. Molecular identification of Cryptosporidium spp. in animal and human hosts from the Czech Republic. Vet. Parasitol. 122, 183–192. Hallier-Soulier, S. and Guillot, E. 2000. Detection of cryptosporidia and Cryptosporidium parvum oocysts in environmental water samples by immunomagnetic separation—polymerase chain reaction. J. Appl. Microbiol. 89, 5–10. Hanninen, M.L., Horman, A., Rimhanen-Finne, R., Vahtera, H., Malmberg, S., Herve, S., and Lahti, K. 2005. Monitoring of Cryptosporidium and Giardia in the Vantaa river basin, southern Finland. Int. J. Hyg. Environ. Health 208, 163–171. Hashimoto, A., Sugimoto, H., Morita, S., and Hirata, T. 2006. Genotyping of single Cryptosporidium oocysts in sewage by semi-nested PCR and direct sequencing. Water Res. 40, 2527–2532. Hirata, T. and Hashimoto, A. 2006. Genotyping of single Cryptosporidium oocysts isolated from sewage and river water. Water Sci. Technol. 54, 197–202. 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. Huang, K., Akiyoshi, D.E., Feng, X., and Tzipori, S. 2003. Development of patent infection in immunosuppressed C57Bl/6 mice with a single Cryptosporidium meleagridis oocyst. J. Parasitol. 89, 620–622. Hunter, P.R., Chalmers, R.M., Syed, Q., Hughes, L.S., Woodhouse, S., and Swift, L. 2003. Foot and mouth disease and cryptosporidiosis: Possible interaction between two emerging infectious diseases. Emerg. Infect. Dis. 9, 109–112. Hunter, P.R., Hughes, S., Woodhouse, S., Raj, N., Syed, Q., Chalmers, R.M., Verlander, N.Q., and Goodacre, J. 2004a. Health sequelae of human cryptosporidiosis in immunocompetent patients. Clin. Infect. Dis. 39, 504–510. Hunter, P.R., Hughes, S., Woodhouse, S., Syed, Q., Verlander, N.Q., Chalmers, R.M., Morgan, K., Nichols, G., Beeching, N., and Osborn, K. 2004b. Sporadic cryptosporidiosis case-control study with genotyping. Emerg. Infect. Dis. 10, 1241–1249. Hunter, P.R. and Nichols, G. 2002. Epidemiology and clinical features of Cryptosporidium infection in immunocompromised patients. Clin. Microbiol. Rev. 15, 145–154. 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. Jellison, K.L., Hemond, H.F., and Schauer, D.B. 2002. Sources and species of Cryptosporidium oocysts in the Wachusett reservoir watershed. Appl. Environ. Microbiol. 68, 569–575. Jiang, J., Alderisio, K.A., Singh, A., and Xiao, L. 2005a. Development of procedures for direct extraction of Cryptosporidium DNA from water concentrates and for relief of PCR inhibitors. Appl. Environ. Microbiol. 71, 1135–1141. Jiang, J., Alderisio, K.A., and Xiao, L. 2005b. Distribution of Cryptosporidium genotypes in storm event water samples from three watersheds in New York. Appl. Environ. Microbiol. 71, 4446–4454. Jiang, J. and Xiao, L. 2003. An evaluation of molecular diagnostic tools for the detection and differentiation of human-pathogenic Cryptosporidium spp. J. Eukaryot. Microbiol. 50 Suppl., 542–547. 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 oocysts in water samples. Appl. Environ. Microbiol. 61, 3849–3855. Karasudani, T., Aoki, S., Takeuchi, J., Okuyama, M., Oseto, M., Matsuura, S., Asai, T., and Inouye, H. 2001. Sensitive detection of Cryptosporidium oocysts in environmental water samples by reverse transcriptionPCR. Jpn. J. Infect. Dis. 54, 122–124. Katsumata, T., Hosea, D., Wasito, E.B., Kohno, S., Hara, K., Soeparto, P., and Ranuh, I.G. 1998. Cryptosporidiosis in Indonesia: A hospital-based study and a community-based survey. Am. J. Trop. Med. Hyg. 59, 628–632.


156


Katsumata, T., Hosea, D., Ranuh, I.G., Uga, S., Yanagi, T., and Kohno, S., 2000. Short report: Possible Cryptosporidium muris infection in humans. Am. J. Trop. Med. Hyg. 62, 70–72. Kaucner, C. and Stinear, T. 1998. Sensitive and rapid detection of viable Giardia cysts and Cryptosporidium parvum oocysts in large-volume water samples with wound fiberglass cartridge filters and reverse transcription-PCR. Appl. Environ. Microbiol. 64, 1743–1749. Kiang, K.M., Scheftel, J.M., Leano, F.T., Taylor, C.M., Belle-Isle, P.A., Cebelinski, E.A., Danila, R., and Smith, K.E. 2006. Recurrent outbreaks of cryptosporidiosis associated with calves among students at an educational farm programme, Minnesota, 2003. Epidemiol. Infect. 134, 878–886. Kimbell, L.M., Miller, D.L., Chavez, W., and Altman, N. 1999. Molecular analysis of the 18S rRNA gene of Cryptosporidium serpentis in a wild-caught corn snake (Elaphe guttata guttata) and a five-species restriction fragment length polymorphism-based assay that can additionally discern C. parvum from C. wrairi. Appl. Environ. Microbiol. 65, 5345–5349. Koudela, B. and Modry, D. 1998. New species of Cryptosporidium (Apicomplexa, Cryptosporidiidae) from lizards. Fol. Parasitol. 45, 93–100. Langkjaer, R.B., Vigre, H., Enemark, H.L., and Maddox-Hyttel, C. 2007. Molecular and phylogenetic characterization of Cryptosporidium and Giardia from pigs and cattle in Denmark. Parasitology (in press). Laxer, M.A., Timblin, B.K., and Patel, R.J. 1991. DNA sequences for the specific detection of Cryptosporidium parvum by the polymerase chain reaction. Am. J. Trop. Med. Hyg. 45, 688–694. Le Blancq, S.M., Khramtsov, N.V., Zamani, F., Upton, S.J., and Wu, T.W. 1997. Ribosomal RNA gene organization in Cryptosporidium parvum. Mol. Biochem. Parasitol. 90, 463–478. Learmonth, J.J., Ionas, G., Ebbett, K.A., and Kwan, E.S. 2004. Genetic characterization and transmission cycles of Cryptosporidium species isolated from humans in New Zealand. Appl. Environ. Microbiol. 70, 3973–3978. Learmonth, J.J., Ionas, G., Pita, A.B., and Cowie, R.S. 2003. Identification and genetic characterisation of Giardia and Cryptosporidium strains in humans and dairy cattle in the Waikato Region of New Zealand. Water Sci. Technol. 47, 21–26. Leav, B.A., Mackay, M.R., Anyanwu, A., RM, O.C., Cevallos, A.M., Kindra, G., Rollins, N.C., Bennish, M.L., Nelson, R.G., and Ward, H.D. 2002. Analysis of sequence diversity at the highly polymorphic Cpgp40/15 locus among Cryptosporidium isolates from human immunodeficiency virus-infected children in South Africa. Infect. Immun. 70, 3881–3890. LeChevallier, M.W., Di Giovanni, 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 spp. in source waters. Appl. Environ. Microbiol. 69, 971–979. Lehmann, T., Graham, D.H., Dahl, E.R., Bahia-Oliveira, L.M., Gennari, S.M., and Dubey, J.P. 2004. Variation in the structure of Toxoplasma gondii and the roles of selfing, drift, and epistatic selection in maintaining linkage disequilibria. Infect. Genet. Evol. 4, 107–114. Leng, X., Mosier, D.A., and Oberst, R.D. 1996. Differentiation of Cryptosporidium parvum, C. muris, and C. baileyi by PCR-RFLP analysis of the 18S rRNA gene. Vet. Parasitol. 62, 1–7. Leoni, F., Amar, C., Nichols, G., Pedraza-Diaz, S., and McLauchlin, J. 2006a. Genetic analysis of Cryptosporidium from 2414 humans with diarrhea in England between 1985 and 2000. J. Med. Microbiol. 55, 703–707. Leoni, F., Gallimore, C.I., Green, J., and McLauchlin, J. 2003a. 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., Gallimore, C.I., Green, J., and McLauchlin, J. 2003b. A rapid method for identifying diversity within PCR amplicons using a heteroduplex mobility assay and synthetic polynucleotides: Application to characterisation of dsRNA elements associated with Cryptosporidium. J. Microbiol. Methods. 54, 95–103. Leoni, F., Gallimore, C.I., Green, J., and McLauchlin, J. 2006b. Characterization of small double stranded RNA molecule in Cryptosporidium hominis, Cryptosporidium felis and Cryptosporidium meleagridis. Parasitol. Int. 55, 299–306. Lindquist, H.D., Bennett, J.W., Ware, M., Stetler, R.E., Gauci, M., and Schaefer, F.W. 2001. Testing methods for detection of Cryptosporidium spp in water samples. Southeast Asian J. Trop. Med. Public Health. 32, 190–194.



157

Lowery, C.J., Millar, B.C., Moore, J.E., Xu, J., Xiao, L., Rooney, P.J., Crothers, L., and Dooley, J.S. 2001a. Molecular genotyping of human cryptosporidiosis in Northern Ireland: Epidemiological aspects and review. Ir. J. Med. Sci. 170, 246–250. Lowery, C.J., Moore, J.E., Millar, B.C., Burke, D.P., McCorry, K.A., Crothers, E., and Dooley, J.S. 2000. Detection and speciation of Cryptosporidium spp. in environmental water samples by immunomagnetic separation, PCR and endonuclease restriction. J. Med. Microbiol. 49, 779–785. Lowery, C.J., Moore, J.E., Millar, B.C., McCorry, K.A., Xu, J., Rooney, P.J., and Dooley, J.S. 2001b. Occurrence and molecular genotyping of Cryptosporidium spp. in surface waters in Northern Ireland. J. Appl. Microbiol. 91, 774–779. Lowery, C.J., Nugent, P., Moore, J.E., Millar, B.C., Xiru, X., and Dooley, J.S. 2001c. PCR-IMS detection and molecular typing of Cryptosporidium parvum recovered from a recreational river source and an associated mussel (Mytilus edulis) bed in Northern Ireland. Epidemiol. Infect. 127, 545–553. Maggi, P., Larocca, A.M., Quarto, M., Serio, G., Brandonisio, O., Angarano, G., and Pastore, G. 2000. Effect of antiretroviral therapy on cryptosporidiosis and microsporidiosis in patients infected with human immunodeficiency virus type 1. Eur. J. Clin. Microbiol. Infect. Dis. 19, 213–217. Mallon, M., MacLeod, A., Wastling, J., Smith, H., Reilly, B., and Tait, A. 2003a. Population structures and the role of genetic exchange in the zoonotic pathogen Cryptosporidium parvum. J. Mol. Evol. 56, 407–417. Mallon, M.E., MacLeod, A., Wastling, J.M., Smith, H., and Tait, A. 2003b. Multilocus genotyping of Cryptosporidium parvum Type 2: Population genetics and sub-structuring. Infect. Genet. Evol. 3, 207–218. Masago, Y., Oguma, K., Katayama, H., and Ohgaki, S. 2006. Quantification and genotyping of Cryptosporidium spp. in river water by quenching probe PCR and denaturing gradient gel electrophoresis. Water Sci. Technol. 54, 119–126. Matsubayashi, M., Abe, N., Takami, K., Kimata, I., Iseki, M., Nakanishi, T., Tani, H., Sasai, K., and Baba, E. 2004. First record of Cryptosporidium infection in a raccoon dog (Nyctereutes procyonoides viverrinus). Vet. Parasitol. 120, 171–175. Mayer, C.L. and Palmer, C.J. 1996. Evaluation of PCR, nested PCR, and fluorescent antibodies for detection of Giardia and Cryptosporidium species in wastewater. Appl. Environ. Microbiol. 62, 2081–2085. 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. McLauchlin, J., Pedraza-Diaz, S., Amar-Hoetzeneder, C., and Nichols, G.L. 1999. Genetic characterization of Cryptosporidium strains from 218 patients with diarrhea diagnosed as having sporadic cryptosporidiosis. J. Clin. Microbiol. 37, 3153–3158. Meamar, A.R., Guyot, K., Certad, G., Dei-Cas, E., Mohraz, M., Mohebali, M., Mohammad, K., Mehbod, A.A., Rezaie, S., and Rezaian, M. 2007. Molecular characterization of Cryptosporidium isolates from humans and animals in Iran. Appl. Environ. Microbiol. 73, 1033–1035. Millard, P.S., Gensheimer, K.F., Addiss, D.G., Sosin, D.M., Beckett, G.A., Houck-Jankoski, A., and Hudson, A. 1994. An outbreak of cryptosporidiosis from fresh-pressed apple cider. J. Am. Med. Assoc. 272, 1592–1596. Misic, Z. and Abe, N. 2007. Subtype analysis of Cryptosporidium parvum isolates from calves on farms around Belgrade, Serbia and Montenegro, using the 60 kDa glycoprotein gene sequences. Parasitology (in press). Molbak, K., Aaby, P., Hojlyng, N., and da Silva, A.P. 1994. Risk factors for Cryptosporidium diarrhea in early childhood: A case-control study from Guinea-Bissau, West Africa. Am. J. Epidemiol. 139, 734–740. 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. Morgan, U., Weber, R., Xiao, L., Sulaiman, I., Thompson, R.C., 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., Xiao, L., Sulaiman, I., Weber, R., Lal, A.A., Thompson, R.C., and Deplazes, P. 1999a. Which genotypes/species of Cryptosporidium are humans susceptible to? J. Eukaryot. Microbiol. 46, 42S and 43S.


158


Morgan, U.M., Constantine, C.C., P, O.D., Meloni, B.P., PA, O.B., and Thompson, R.C. 1995. Molecular characterization of Cryptosporidium isolates from humans and other animals using random amplified polymorphic DNA analysis. Am. J. Trop. Med. Hyg. 52, 559–564. Morgan, U.M., Deplazes, P., Forbes, D.A., Spano, F., Hertzberg, H., Sargent, K.D., Elliot, A., and Thompson, R.C. 1999b. Sequence and PCR-RFLP analysis of the internal transcribed spacers of the rDNA repeat unit in isolates of Cryptosporidium from different hosts. Parasitology 118, 49–58. Morgan, U.M., Monis, P.T., Fayer, R., Deplazes, P., and Thompson, R.C. 1999c. Phylogenetic relationships among isolates of Cryptosporidium: Evidence for several new species. J. Parasitol. 85, 1126–1133. Morgan, U.M., Pallant, L., Dwyer, B.W., Forbes, D.A., Rich, G., and Thompson, R.C. 1998. Comparison of PCR and microscopy for detection of Cryptosporidium parvum in human fecal specimens: Clinical trial. J. Clin. Microbiol. 36, 995–998. Morgan, U.M., Sturdee, A.P., Singleton, G., Gomez, M.S., Gracenea, M., Torres, J., Hamilton, S.G., Woodside, D.P., and Thompson, R.C.A. 1999d. 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. 2000b. Epidemiology and strain variation of Cryptosporidium parvum. Contrib. Microbiol. 6, 116–139. Morgan, U.M., Xiao, L., Hill, B.D., O’Donoghue, P., Limor, J., Lal, A., and Thompson, R.C. 2000c. Detection of the Cryptosporidium parvum “human” genotype in a dugong (Dugong dugon). J. Parasitol. 86, 1352–1354. 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. Navarro-i-Martinez, L., da Silva, A.J., Garces, G., Montoya Palacio, M.N., del Aguila, C., and Bornay-Llinares, F.J. 2006. Cryptosporidiosis in HIV-positive patients from Medellin, Colombia. J. Eukaryot. Microbiol. 53 Suppl., S37–S39. Neira-Otero, P., Munoz-Saldias, N., Sanchez-Moreno, M., and Rosales-Lombardo, M.J. 2005. Molecular characterization of Cryptosporidium species and genotypes in Chile. Parasitol. Res. 97, 63–67. Ngouanesavanh, T., Guyot, K., Certad, G., Fichoux, Y.L., Chartier, C., Verdier, R.-I., Cailliez, J.-C., Camus, D., Dei-Cas, E., and Banuls, A.-L. 2006. Cryptosporidium population genetics: Evidence of clonality in isolates from France and Haiti. J. Eukaryot. Microbiol. 53, S33–S36. Nichols, R.A., Campbell, B.M., and Smith, H.V. 2003. Identification of Cryptosporidium spp. oocysts in United Kingdom noncarbonated natural mineral waters and drinking waters by using a modified nested PCR-restriction fragment length polymorphism assay. Appl. Environ. Microbiol. 69, 4183–4189. Nichols, R.A., Campbell, B.M., and Smith, H.V. 2006. Molecular fingerprinting of Cryptosporidium oocysts isolated during water monitoring. Appl. Environ. Microbiol. 72, 5428–5435. Nichols, G.L., Chalmers, R.M., Sopwith, W., Regan, M., Hunter, C.A., Grenfell, P., Harrison, F., and Lane, C. 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, U.K. (http://www.fwr.org/waterq/dwi0849.htm as of January 1, 2007). Ong, C.S., Eisler, D.L., Alikhani, A., Fung, V.W., Tomblin, J., Bowie, W.R., and Isaac-Renton, J.L. 2002. Novel Cryptosporidium genotypes in sporadic cryptosporidiosis cases: First report of human infections with a cervine genotype. Emerg. Infect. Dis. 8, 263–268. Ong, C.S., Eisler, D.L., Goh, S.H., Tomblin, J., Awad-El-Kariem, F.M., Beard, C.B., Xiao, L., Sulaiman, I., Lal, A., Fyfe, M., King, A., Bowie, W.R., and Isaac-Renton, J.L. 1999. Molecular epidemiology of cryptosporidiosis outbreaks and transmission in British Columbia, Canada. Am. J. Trop. Med. Hyg. 61, 63–69. Ong, C.S.L., Chow, S., So, P.P.L., Chen, R., Xiao, L., Sulaiman, I., Zhou, L., Ellis, A., Aramini, J., Horsman, G., and Isaac-Renton, J.L. 2005. Identification of two different Cryptosporidium hominis subtypes from cases in the 2001 waterborne cryptosporidiosis outbreak in North Battleford, Saskatchewan, in Proceedings of the 11th Canadian National Conference and 2nd Policy Forum on Drinking Water (Calgary, Canada, April 3–6, 2004), Canadian Water and Wastewater Association, Ottawa, Ontario, pp. 628–638. Ortega-Mora, L.M., Requejo-Fernandez, J.A., Pilar-Izquierdo, M., and Pereira-Bueno, J. 1999. Role of adult sheep in transmission of infection by Cryptosporidium parvum to lambs: Confirmation of periparturient rise. Int. J. Parasitol. 29, 1261–1268.



159

Oura, C.A., Asiimwe, B.B., Weir, W., Lubega, G.W., and Tait, A. 2005. Population genetic analysis and substructuring of Theileria parva in Uganda. Mol. Biochem. Parasitol. 140, 229–239. Palmer, C.J., Xiao, L., Terashima, A., Guerra, H., Gotuzzo, E., Saldias, 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. Park, J.H., Guk, S.M., Han, E.T., Shin, E.H., Kim, J.L., and Chai, J.Y. 2006. Genotype analysis of Cryptosporidium spp. prevalent in a rural village in Hwasun-gun, Republic of Korea. Korean J. Parasitol. 44, 27–33. 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. Pedraza-Diaz, S., Amar, C., Iversen, A.M., Stanley, P.J., and McLauchlin, J. 2001a. Unusual Cryptosporidium species recovered from human faeces: First description of Cryptosporidium felis and Cryptosporidium “dog type” from patients in England. J. Med. Microbiol. 50, 293–296. Pedraza-Diaz, S., Amar, C., and McLauchlin, J. 2000. The identification and characterisation of an unusual genotype of Cryptosporidium from human faeces as Cryptosporidium meleagridis. FEMS Microbiol. Lett. 189, 189–194. Pedraza-Diaz, S., Amar, C., Nichols, G.L., and McLauchlin, J. 2001b. Nested polymerase chain reaction for amplification of the Cryptosporidium oocyst wall protein gene. Emerg. Infect. Dis. 7, 49–56. Pedraza-Diaz, S., Amar, C.F., 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 Youngs, E.R. 2001c. Cryptosporidium meleagridis from humans: Molecular analysis and description of affected patients. J. Infect. 42, 243–250. Peng, M.M., Matos, O., Gatei, W., Das, P., Stantic-Pavlinic, M., Bern, C., Sulaiman, I.M., Glaberman, S., Lal, A.A., and Xiao, L. 2001. A comparison of Cryptosporidium subgenotypes from several geographic regions. J. Eukaryot. Microbiol. 48, Suppl., 28S–31S. Peng, M.M., Meshnick, S.R., Cunliffe, N.A., Thindwa, B.D., Hart, C.A., Broadhead, R.L., and Xiao, L. 2003a. Molecular epidemiology of cryptosporidiosis in children in Malawi. J. Eukaryot. Microbiol. 50, Suppl., 557–559. Peng, M.M., Wilson, M.L., Holland, R.E., Meshnick, S.R., Lal, A.A., and Xiao, L. 2003b. Genetic diversity of Cryptosporidium spp. in cattle in Michigan: Implications for understanding the transmission dynamics. Parasitol. Res. 90, 175–180. Peng, M.M., Xiao, L., Freeman, A.R., Arrowood, M.J., Escalante, A.A., Weltman, A.C., Ong, C.S., Mac Kenzie, W.R., Lal, A.A., and Beard, C.B. 1997. Genetic polymorphism among Cryptosporidium parvum isolates: Evidence of two distinct human transmission cycles. Emerg. Infect. Dis. 3, 567–573. Pereira, S.J., Ramirez, N.E., Xiao, L., and Ward, L.A. 2002. Pathogenesis of human and bovine Cryptosporidium parvum in gnotobiotic pigs. J. Infect. Dis. 186, 715–718. 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. Pieniazek, N.J., Bornay-Llinares, F.J., Slemenda, S.B., da Silva, A.J., Moura, I.N., Arrowood, M.J., Ditrich, O., and Addiss, D.G. 1999. New Cryptosporidium genotypes in HIV-infected persons. Emerg. Infect. Dis. 5, 444–449. 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. Preiser, G., Preiser, L., and Madeo, L. 2003. An outbreak of cryptosporidiosis among veterinary science students who work with calves. J. Am. Coll. Health. 51, 213–215. Priest, J.W., Kwon, J.P., Arrowood, M.J., and Lammie, P.J. 2000. Cloning of the immunodominant 17-kDa antigen from Cryptosporidium parvum. Mol. Biochem. Parasitol. 106, 261–271. Quiroz, E.S., Bern, C., MacArthur, J.R., Xiao, L., Fletcher, M., Arrowood, M.J., Shay, D.K., Levy, M.E., Glass, R.I., and Lal, A. 2000. An outbreak of cryptosporidiosis linked to a foodhandler. J. Infect. Dis. 181, 695–700. 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.


160


Robertson, B., Sinclair, M.I., Forbes, A.B., Veitch, M., Kirk, M., Cunliffe, D., Willis, J., and Fairley, C.K. 2002. Case-control studies of sporadic cryptosporidiosis in Melbourne and Adelaide, Australia. Epidemiol. Infect. 128, 419–431. Robinson, R.A. and Pugh, R.N. 2002. Dogs, zoonoses and immunosuppression. J. R. Soc. Health. 122, 95–98. Rochelle, P.A., De Leon, R., Stewart, M.H., and Wolfe, R.L. 1997. Comparison of primers and optimization of PCR conditions for detection of Cryptosporidium parvum and Giardia lamblia in water. Appl. Environ. Microbiol. 63, 106–114. Roy, S.L., DeLong, S.M., Stenzel, S.A., Shiferaw, B., Roberts, J.M., Khalakdina, A., Marcus, R., Segler, S.D., Shah, D.D., Thomas, S., Vugia, D.J., Zansky, S.M., Dietz, V., and Beach, M.J. 2004. Risk factors for sporadic cryptosporidiosis among immunocompetent persons in the United States from 1999 to 2001. J. Clin. Microbiol. 42, 2944–2951. Ruecker, N.J., Bounsombath, N., Wallis, P., Ong, C.S., Isaac-Renton, J.L., and Neumann, N.F. 2005. Molecular forensic profiling of Cryptosporidium species and genotypes in raw water. Appl. Environ. Microbiol. 71, 8991–8994. Ruecker, N.J., Braithwaite, S.L., Topp, E., Edge, T., Lapen, D.R., Wilkes, G., Robertson, W., Medeiros, D., Sensen, C.W., and Neumann, N.F. 2007. Tracking host sources of Cryptosporidium spp. in raw water for improved health risk assessment. Appl. Environ. Microbiol. 73, 3945–3957. Ryan, U., Read, C., Hawkins, P., Warnecke, M., Swanson, P., Griffith, M., Deere, D., Cunningham, M., and Cox, P. 2005a. Genotypes of Cryptosporidium from Sydney water catchment areas. J. Appl. Microbiol. 98, 1221–1229. 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., Bath, C., Robertson, I., Read, C., Elliot, A., McInnes, L., Traub, R., and Besier, B. 2005b. Sheep may not be an important zoonotic reservoir for Cryptosporidium and Giardia parasites. Appl. Environ. Microbiol. 71, 4992–4997. 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. Santín, M., Dixon, B.R., and Fayer, R. 2005. Genetic characterization of Cryptosporidium isolates from ringed seals (Phoca hispida) in Northern Quebec, Canada. J. Parasitol. 91, 712–716. Santín, M. and Fayer, R. 2007a. Intragenotypic variations in the Cryptosporidium cervine genotype from sheep with implications for public health. J. Parasitol. 93, 668–672. 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. Santín, M., Trout, J.M., and Fayer, R. 2007b. Prevalence and molecular characterization of Cryptosporidium and Giardia species and genotypes in sheep in Maryland. Vet. Parasitol. 146, 17–24. Schindler, A.R., Abs El-Osta, Y.G., Stevens, M., Sinclair, M.I., and Gasser, R.B. 2005. Capillary electrophoretic analysis of fragment length polymorphism in ribosomal markers of Cryptosporidium from humans. Mol. Cell. Probes. 19, 394–399. Sinclair, J.L. 2000. Enumeration of Cryptosporidium spp. in water with US EPA method 1622, USA. J. AOAC Int. 83, 1108–1114. Skerrett, H.E. and Holland, C.V. 2001. Asymptomatic shedding of Cryptosporidium oocysts by red deer hinds and calves. Vet. Parasitol. 94, 239–246. Slapeta, J. 2006. Cryptosporidium species found in cattle: A proposal for a new species. Trends Parasitol., 22, 469–474. Sluter, S.D., Tzipori, S., and Widmer, G. 1997. Parameters affecting polymerase chain reaction detection of waterborne Cryptosporidium parvum oocysts. Appl. Microbiol. Biotechnol. 48, 325–330. Smerdon, W.J., Nichols, T., Chalmers, R.M., Heine, H., and Reacher, M.H. 2003. Foot and mouth disease in livestock and reduced cryptosporidiosis in humans, England and Wales. Emerg. Infect. Dis. 9, 22–28. Smith, A., Reacher, M., Smerdon, W., Adak, G.K., Nichols, G., and Chalmers, R.M. 2006a. Outbreaks of waterborne infectious intestinal disease in England and Wales, 1992–2003. Epidemiol. Infect. 1–9. Smith, H.V., Caccio, S.M., Tait, A., McLauchlin, J., and Thompson, R.C. 2006b. Tools for investigating the environmental transmission of Cryptosporidium and Giardia infections in humans. Trends Parasitol. 22, 160–167.



161

Smith, H.V., Nichols, R.A., Mallon, M., Macleod, A., Tait, A., Reilly, W.J., Browning, L.M., Gray, D., Reid, S.W., and Wastling, J.M. 2005. Natural Cryptosporidium hominis infections in Scottish cattle. Vet. Rec. 156, 710 and 711. Smith, K.E., Stenzel, S.A., Bender, J.B., Wagstrom, E., Soderlund, D., Leano, F.T., Taylor, C.M., Belle-Isle, P.A., and Danila, R. 2004. Outbreaks of enteric infections caused by multiple pathogens associated with calves at a farm day camp. Pediatr. Infect. Dis. J. 23, 1098–1104. Soba, B., Petrovec, M., Mioc, V., and Logar, J. 2006. Molecular characterisation of Cryptosporidium isolates from humans in Slovenia. Clin. Microbiol. Infect. 12, 918–921. Sopwith, W., Osborn, K., Chalmers, R., and Regan, M. 2005. The changing epidemiology of cryptosporidiosis in North West England. Epidemiol. Infect. 133, 785–793. Strong, W.B., Gut, J., and Nelson, R.G. 2000. Cloning and sequence analysis of a highly polymorphic Cryptosporidium parvum gene encoding a 60-kilodalton glycoprotein and characterization of its 15- and 45-kilodalton zoite surface antigen products. Infect. Immun. 68, 4117–4134. 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. Sturbaum, G.D., Klonicki, P.T., Marshall, M.M., Jost, B.H., Clay, B.L., and Sterling, C.R. 2002. Immunomagnetic separation (IMS)-fluorescent antibody detection and IMS-PCR detection of seeded Cryptosporidium parvum oocysts in natural waters and their limitations. Appl. Environ. Microbiol. 68, 2991–2996. Sturbaum, G.D., Reed, C., Hoover, P.J., Jost, B.H., Marshall, M.M., and Sterling, C.R. 2001. Species-specific, nested PCR-restriction fragment length polymorphism detection of single Cryptosporidium parvum oocysts. Appl. Environ. Microbiol. 67, 2665–2668. Sulaiman, I.M., Hira, P.R., Zhou, L., Al-Ali, F.M., Al-Shelahi, F.A., Shweiki, H.M., Iqbal, J., Khalid, N., and Xiao, L. 2005. Unique endemicity of cryptosporidiosis in children in Kuwait. J. Clin. Microbiol. 43, 2805–2809. Sulaiman, I.M., Lal, A.A., and Xiao, L. 2001. A population genetic study of the Cryptosporidium parvum human genotype parasites. J. Eukaryot. Microbiol. 48 Suppl., 24S–27S. Sulaiman, I.M., Lal, A.A., and Xiao, L. 2002. Molecular phylogeny and evolutionary relationships of Cryptosporidium parasites at the actin locus. J. Parasitol. 88, 388–394. Sulaiman, I.M., Morgan, U.M., Thompson, R.C., Lal, A.A., and Xiao, L. 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., Xiao, L., and Lal, A.A. 1999. Evaluation of Cryptosporidium parvum genotyping techniques. Appl. Environ. Microbiol. 65, 4431–4435. Sulaiman, I.M., Xiao, L., Yang, C., Escalante, L., Moore, A., Beard, C.B., Arrowood, M.J., and Lal, A.A. 1998. Differentiating human from animal isolates of Cryptosporidium parvum. Emerg. Infect. Dis. 4, 681–685. Sunnotel, O., Snelling, W.J., Xiao, L., Moule, K., Moore, J.E., Millar, B.C., Dooley, J.S., and Lowery, C.J. 2006. Rapid and sensitive detection of single Cryptosporidium oocysts from archived glass slides. J. Clin. Microbiol. 44, 3285–3291. Tait, A., Wastling, J.M., Smith, H., MacLeod, A., and Mallon, M.E. 2004. Response to: Population genetics of Cryptosporidium parvum. Trends Parasitol. 20, 6. Tanriverdi, S., Arslan, M.O., Akiyoshi, D.E., Tzipori, S., and Widmer, G. 2003. Identification of genotypically mixed Cryptosporidium parvum populations in humans and calves. Mol. Biochem. Parasitol. 130, 13–22. Tanriverdi, S., Markovics, A., Arslan, M.O., Itik, A., Shkap, V., and Widmer, G. 2006. Emergence of distinct genotypes of Cryptosporidium parvum in structured host populations. Appl. Environ. Microbiol. 72, 2507–2513. Tanriverdi, S. and Widmer, G. 2006. Differential evolution of repetitive sequences in Cryptosporidium parvum and Cryptosporidium hominis. Infect. Genet. Evol. 6, 113–122. Thompson, H.P., Dooley, J.S., Kenny, J., McCoy, M., Lowery, C.J., Moore, J.E., and Xiao, L. 2007. Genotypes and subtypes of Cryptosporidium spp. in neonatal calves in Northern Ireland. Parasitol. Res. 100, 619–624. Tiangtip, R. and Jongwutiwes, S. 2002. Molecular analysis of Cryptosporidium species isolated from HIVinfected patients in Thailand. Trop. Med. Int. Health. 7, 357–364. Tibayrenc, M. and Ayala, F. 2002. The clonal theory of parasitic protozoa: 12 years on. Trends Parasitol. 18, 405.


162


Trotz-Williams, L.A., Martin, D.S., Gatei, W., Cama, V., Peregrine, A.S., Martin, S.W., Nydam, D.V., Jamieson, F., and Xiao, L. 2006. Genotype and subtype analyses of Cryptosporidium isolates from dairy calves and humans in Ontario. Parasitol. Res. 99, 346–352. Trout, J.M., Santín, M., Fayer, R., 2006. Giardia and Cryptosporidium species and genotypes in coyotes (Canis latrans). J. Zoo Wildl. Med. 37, 141–144. Tsushima, Y., Karanis, P., Kamada, T., Xuan, X., Makala, L.H., Tohya, Y., Akashi, H., and Nagasawa, H. 2003. Viability and infectivity of Cryptosporidium parvum oocysts detected in river water in Hokkaido, Japan. J. Vet. Med. Sci. 65, 585–589. 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. 2003. Cryptosporidium parvum in children with diarrhea in Mulago Hospital, Kampala, Uganda. Am. J. Trop. Med. Hyg. 68, 710–715. Tzipori, S. and Griffiths, J.K. 1998. Natural history and biology of Cryptosporidium parvum. Adv. Parasitol. 40, 5–36. 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. Weintraub, J.M. 2006. Improving Cryptosporidium testing methods: A public health perspective. J. Water. Health. 4 Suppl., 1, 23–26. Widmer, G. 2004. Population genetics of Cryptosporidium parvum. Trends Parasitol. 20, 3–6. 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. Widmer, G., Feng, X., and Tanriverdi, S. 2004. Genotyping of Cryptosporidium parvum with microsatellite markers. Methods Mol. Biol. 268, 177–188. 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. Winter, G., Gooley, A.A., Williams, K.L., and Slade, M.B. 2000. Characterization of a major sporozoite surface glycoprotein of Cryptosporidium parvum. Funct. Integr. Genomics. 1, 207–217. 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 RFLPsingle-strand conformational polymorphism analyses. Appl. Environ. Microbiol. 69, 4720–4726. 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. Xiao, L., Alderisio, K., and Singh, A. 2006a. Development and Standardization of a Cryptosporidium Genotyping Tool for Water Samples. Awwa Research Foundation, Denver, CO., pp. 116. Xiao, L., Alderisio, K.A., and Jiang, J. 2006b. 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., Arrowood, M., Sulaiman, I., Zhou, L., Kawai, V., Vivar, A., Lal, A.A., and Gilman, R.H. 2002a. Identification of the Cryptosporidium pig genotype in a human patient. J. Infect. Dis. 185, 1846–1848. 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., Bern, C., Sulaiman, I.M., and Lal, A.A. 2004a. Molecular epidemiology of human cryptosporidiosis, in Cryptosporidium: From Molecules to Disease, Thompson, R.C.A., Armson, A., and Ryan, U.M., Eds., Elsevier, Amsterdam, pp. 121–146. Xiao, L., Escalante, L., Yang, C., Sulaiman, I., Escalante, A.A., Montali, R.J., Fayer, R., and Lal, A.A. 1999a. 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. 2004b. Cryptosporidium taxonomy: Recent advances and implications for public health. Clin. Microbiol. Rev. 17, 72–97.



163

Xiao, L., Fayer, R., Ryan, U.M., and Upton, S.J. 2007a. Response to the newly proposed species Cryptosporidium pestis. Trends Parasitol., 23, 41–42. Xiao, L., Herd, R.P., and McClure, K.E. 1994. Periparturient rise in the excretion of Giardia sp. cysts and Cryptosporidium parvum oocysts as a source of infection for lambs. J. Parasitol. 80, 55–59. Xiao, L., Lal, A.A., and Jiang, J. 2004c. Detection and differentiation of Cryptosporidium oocysts in water by PCR-RFLP. Methods 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., Limor, J.R., Li, L., Morgan, U., Thompson, R.C., and Lal, A.A. 1999b. Presence of heterogeneous copies of the small subunit rRNA gene in Cryptosporidium parvum human and marsupial genotypes and Cryptosporidium felis. J. Eukaryot. Microbiol. 46, 44S and 45S. Xiao, L., Morgan, U.M., Limor, J., Escalante, A., Arrowood, M., Shulaw, W., Thompson, R.C., Fayer, R., and Lal, A.A. 1999c. Genetic diversity within Cryptosporidium parvum and related Cryptosporidium species. Appl. Environ. Microbiol. 65, 3386–3391. Xiao, L., and Ryan, U.M. 2004. Cryptosporidiosis: An update in molecular epidemiology. Curr. Opin. Infect. Dis. 17, 483–490. Xiao, L., Ryan, U.M., Graczyk, T.K., Limor, J., Li, L., Kombert, M., Junge, R., Sulaiman, I.M., Zhou, L., Arrowood, M.J., Koudela, B., Modry, D., and Lal, A.A. 2004d. Genetic diversity of Cryptosporidium spp. in captive reptiles. Appl. Environ. Microbiol. 70, 891–899. Xiao, L., Singh, A., Limor, J., Graczyk, T.K., Gradus, S., and Lal, A. 2001c. Molecular characterization of Cryptosporidium oocysts in samples of raw surface water and wastewater. Appl. Environ. Microbiol. 67, 1097–1101. Xiao, L., Sulaiman, I.M., Ryan, U.M., Zhou, L., Atwill, E.R., Tischler, M.L., Zhang, X., Fayer, R., and Lal, A.A. 2002b. Host adaptation and host-parasite co-evolution in Cryptosporidium: Implications for taxonomy and public health. Int. J. Parasitol. 32, 1773–1785. Xiao, L., Zhou, L., Santín, M., Yang, W., and Fayer, R. 2007b. Distribution of Cryptosporidium parvum subtypes in calves in eastern United States. Parasitol. Res. 100, 701–706. 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. 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., Fayer, R., Trout, J.M., Ryan, U.M., Schaefer, F.W., 3rd and Xiao, L. 2004a. Genotypes of Cryptosporidium species infecting fur-bearing mammals differ from those of species infecting humans. Appl. Environ. Microbiol. 70, 7574–7577. Zhou, L., Kassa, H., Tischler, M.L., and Xiao, L. 2004b. Host-adapted Cryptosporidium spp. in Canada geese (Branta canadensis). Appl. Environ. Microbiol. 70, 4211–4215. 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.


164


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.


Molecular Epidemiology B.

165

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:


166

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



167

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:


168

Cryptosporidium and Cryptosporidiosis, Second Edition 10× Perkin–Elmer PCR buffer dNTP (1.5 mM) GP60-F1 primer (Table A.2) (40 ng/µL) GP60-R1 primer (Table A.2) (40 ng/µL) MgCl2 (25 mM) Nonacetylated bovine serum albumin (10 mg/ML) Distilled water Taq polymerase

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′).



169

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).


170


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



171

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.



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


174


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


Diagnostics

175

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.


176


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.


Diagnostics

177

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)


178


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


Diagnostics

179

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



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


Diagnostics 181


182


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.


Diagnostics

183

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.


184


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.


Diagnostics

185

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).


186


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.


Diagnostics a.

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.


188


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.


Diagnostics

189

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).


190


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


Diagnostics

191

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


192


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.


Diagnostics

193

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.


194


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


Diagnostics

195

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


196


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.


Diagnostics

197

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


198


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 ( 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,


Clinical Disease and Pathology

237

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


238


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



239

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


240


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.



241

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


242


(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



243

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).


244 D.

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.

References Abdel-Messih, I.A., Wierzba, T.F., Abu-Elyazeed, R., Ibrahim, A.F., Ahmed, S.F., Kamal, K., Sanders, J. and Frenck, R. 2005. Diarrhea associated with Cryptosporidium parvum among young children of the Nile River Delta in Egypt. J. Trop. Pediatr. 51, 154–159. Abdo, A., Klassen, J., Urbanski, S., Raber, E. and Swain, M.G. 2003. Reversible sclerosing cholangitis secondary to cryptosporidiosis in a renal transplant patient. J. Hepatol. 38, 688–691. Adal, K.A., Sterling, C.R. and Guerrant R.L. 1995. Cryptosporidium and related species, in Infections of the Gastrointestinal Tract, Blaser, M.J., Smith, P.D., Ravdin, J.I., Greenberg, H.B. and Guerrant, R.L., Eds., New York, Raven Press, 1107–1128. Adams, R.B., Guerrant, R.L., Zu, S., Fang, G. and Roche, J.K. 1994. Cryptosporidium parvum infection of intestinal epithelium: Morphologic and functional studies in an in vitro model. J. Infect. Dis. Jan 169, 170–177. Addy, P.A.K. and Aikins-Bekoe, P. 1986. Cryptosporidiosis in diarrhoeal children in Kumasi, Ghana. Lancet. 1, 735. Adjei, A.A., Armah, H., Rodrigues, O., Renner, L., Borketey, P., Ayeh-Kumi, P., Adiku, T., Sifah, E. and Lartey, M. 2004. Cryptosporidium spp., a frequent cause of diarrhea among children at the Korle-Bu Teaching Hospital, Accra, Ghana. Jpn. J. Infect. Dis. 57, 216–219.



245

Agnew, D.G., Lima, A.A., Newman, R.D., Wuhib, T., Moore, R.D., Guerrant, R.L. and Sears, C.L. 1998. Cryptosporidiosis in northeastern Brazilian children: Association with increased diarrhea morbidity. J. Infect. Dis. 177, 754–760. Alcantara, C.S., Yang, C.H., Steiner, T.S., Barrett, L.J., Lima, A.A., Chappell, C.L., Okhuysen, P.C., White, A.C., Jr and Guerrant, R.L. 2003. Interleukin-8, tumor necrosis factor-alpha, and lactoferrin in immunocompetent hosts with experimental and Brazilian children with acquired cryptosporidiosis. Am. J. Trop. Med. Hyg. 68, 325–328. Amadi, B., Kelly, P., Mwiya, M., Mulwazi, E., Sianongo, S., Changwe, F., Thomson, M., Hachungula, J., Watuka, A., Walker-Smith, J. and Chintu, C. 2001. Intestinal and systemic infection, HIV, and mortality in Zambian children with persistent diarrhea and malnutrition. J. Pediatr. Gastroenterol. Nutr. 32, 550–554. Argenzio, R.A., Lecce, J. and Powell, D.W. 1993. Prostanoids inhibit intestinal NaCl absorption in experimental porcine cryptosporidiosis. Gastroenterology 104, 440–447. Argenzio, R.A., Rhoads, J.M., Armstrong, M. and Gomez G. 1994. Glutamine stimulates prostaglandinsensitive Na+-H+ exchange in experimental porcine cryptosporidiosis. Gastroenterol. 106, 1418–1428. Bannister, P. and Mountford, R.A. Cryptosporidium in the elderly: A cause of life-threatening diarrhea. 1989. Am. J. Med. 86, 507–508. Banwat, E.B., Egah, D.Z., Onile, B.A., Angyo, I.A. and Audu, E.S. 2003. Prevalence of cryptosporidium infection among undernourished children in Jos, Central Nigeria. Niger. Postgrad. Med. J. 10, 84–87. Barbot, L., Topouchian, A., Capet, C., Magne, D., Huneau, J.F., Kapel, N., Gobert, J.G. 2001. Cryptosporidium parvum: Functional study of the intestinal malabsorption syndrome. Ann. Pharm. Fr. 59,305–311. Baxby, D., Hart, C.A. and Blundell, N. 1985. Shedding of oocysts by immunocompetent individuals with cryptosporidiosis. J. Hyg. (Lond). 95, 703–709. Berk, R.N., Wall, S.D., McArdle, C.B., McCutchan ,J.A., Clemett, A.R., Rosenblum, J.S., Premkumer, A. and Megibow, A.J. 1984. Cryptosporidiosis of the stomach and small intestine in patients with AIDS. Am. J. Radiol. 143, 549–554. Bhattacharya, M.K., Teka, T., Faruque, A.S. and Fuchs, G.J. 1997. Cryptosporidium infection in children in urban Bangladesh. J. Trop. Pediatr. 43, 282–286. Blagburn, B.L. and Current, W.L. 1983. Accidental infection of a researcher with human Cryptosporidium. J. Infect. Dis. 148, 772. Blanshard, C., Jackson, A.M., Shanson, D.C., Francis, N. and Gazzard, B.G. 1992. Cryptosporidiosis in HIVseropositive patients. Q. J. Med. 85, 813–823. Buch, K., Nguyen, S., Divino, C.M., Weber, K. and Morotti, R.A. 2005. Cryptosporidiosis presenting as acute appendicitis: A case report. Am. Surg. 71, 537–538. Bushen, O.Y. 2006. Cryptosporidiosis in Guerrant, R.L., Walker, D.H., Weller, P.F. (Eds.) Tropical Diseases: Priniciples, Pathogens, and Practice, Elsevier, Philadelphia, 1003–1014. Bushen, O.Y., Kohli, A., Pinkerton, R.C., Dupnik, K., Newman, R.D., Sears, C.L., Fayer, R., Lima, A.A. and Guerrant, R.L. 2007. Heavy cryptosporidial infections in children in northeast Brazil: Comparison of Cryptosporidium hominis and Cryptosporidium parvum. Trans. R. Soc. Trop. Hyg. 101,378–384. Calzetti, C., Magnani, G., Confalonieri, D., Capelli, A., Moneta, S., Scognamiglio, P. and Fiaccadori, F. 1997. Pancreatitis caused by Cryptosporidium parvum in patients with severe immunodeficiency related to HIV infection. Ann. Ital. Med. Int. 12, 63–66. Campos, M., Jouzdani, E., Sempoux, C., Buts, J.P., Reding, R., Otte, J.B. and Sokal, E.M. 2000. Sclerosing cholangitis associated to cryptosporidiosis in liver-transplanted children. Eur. J. Pediatr. 159, 113–115. Capet, C., Kapel, N., Huneau, J.F., Magne, D., Laikuen, R., Tricottet, V., Benhamou, Y, Tome, D. and Gobert, JG. 1999. Cryptosporidium parvum infection in suckling rats: Impairment of mucosal permeability and Na(+)-glucose cotransport. Exp. Parasitol. 91,119–125. Carvalho-Almeida, T.T., Pinto, P.L., Quadros, C.M., Torres, D.M., Kanamura, H.Y. and Casimiro, A.M. 2006. Detection of Cryptosporidium sp. in non diarrheal faeces from children, in a day care center in the city of Sao Paulo, Brazil. Rev. Inst. Med. Trop. Sao Paulo 48, 27–32. Epub 2006 Mar 9. Casemore, D.P., Sands, R.L. and Curry, A. 1985. Cryptosporidium species: A “new” human pathogen. J. Clin. Pathol. 38:1321–1336. Centers for Disease Control. 1982. Cryptosporidiosis: Assessment of chemotherapy of males with acquired immune deficiency syndrome (AIDS). MMWR. 31, 589.


246


Centers for Disease Control. 1995. Assessing the public health threat associated with waterborne cryptosporidiosis: Report of a workshop. MMWR. 44, 1–19. Cerosimo, E., Wilkowske, C.J., Rosenblatt, J.E. and Ludwig, J. 1992. Isolated antral narrowing associated with gastrointestinal cryptosporidiosis in acquired immunodeficiency syndrome. Mayo Clinic Proc. 67, 553–556. Chai, J.Y., Kim, N.Y., Guk, S.M., Park, Y.K., Seo, M., Han, E.T. and Lee, S.H. 2001. High prevalence and seasonality of cryptosporidiosis in a small rural village occupied predominantly by aged people in the Republic of Korea. Am. J. Trop. Med. Hyg. 65, 518–522. Chaisson, M.A. et al: A prevalence survey for Cryptosporidium and other parasites in healthy homosexual men in New York City. The 34th Annual Meeting of the American Society of Tropical Medicine and Hygiene, November, 1985, abstr 203. Chappell, C.L., Okhuysen, P.C., Langer-Curry, R., Widmer, G., Akiyoshi, D.E., Tanriverdi, S. and Tzipori, S. 2006. Cryptosporidium hominis: Experimental challenge of healthy adults. Am. J. Trop. Med. Hyg. 75,851–857. Chappell, C.L., Okhuysen, P.C., Sterling, C.R. and DuPont, H.L. 1996. Cryptosporidium parvum: Intensity of infection and oocyst excretion patterns in healthy volunteers. J. Infect. Dis. 173,232–236. Chappell, C.L., Okhuysen, P.C., Sterling, C.R., Wang, C., Jakubowski, W. and Dupont, H.L. 1999. Infectivity of Cryptosporidium parvum in healthy adults with pre-existing anti-C. parvum serum immunoglobulin G. Am. J. Trop. Med. Hyg. 60, 157–164. Checkley, W., Gilman, R.H., Epstein, L.D., Suarez, M., Diaz, J.F., Cabrera, L., Black, R.E. and Sterling, C.R. 1997. Asymptomatic and symptomatic cryptosporidiosis: Their acute effect on weight gain in Peruvian children. Am. J. Epidemiol. 145, 156–163. Checkley, W., Epstein, L.D., Gilman, R.H., Black, R.E., Cabrera, L. and Sterling, C.R. 1998. Effects of Cryptosporidium parvum infection in Peruvian children: Growth faltering and subsequent catch-up growth. Am. J. Epidemiol. 148,497–506. Chen XM, Gores GJ, Paya CV, LaRusso NF. 1999. Cryptosporidium parvum induces apoptosis in biliary epithelia by a Fas/Fas ligand-dependent mechanism. Am. J. Physiol. 277,G599–G608. Chen, X.M. and LaRusso, N.F. 2000. Mechanisms of attachment and internalization of Cryptosporidium parvum to biliary and intestinal epithelial cells. Gastroenterology. 118, 368–379. Chen, X.M. and LaRusso, N.F. 2002. Cryptosporidiosis and the pathogenesis of AIDS-cholangiopathy. Semin. Liver Dis. 22, 277–289. Chen, X.M., Levine, S.A., Splinter, P.L., Tietz, P.S., Ganong, A.L., Jobin, C., Gores, G.J., Paya, C.V. and LaRusso, N.F. 2001. Cryptosporidium parvum activates nuclear factor kappaB in biliary epithelia preventing epithelial cell apoptosis. Gastroenterology. 120, 1774–1783. Chen, X.M., Levine, S.A., Tietz, P., Krueger, E., McNiven, M.A., Jefferson, D.M., Mahle, M. and LaRusso NF. 1998. Cryptosporidium parvum is cytopathic for cultured human biliary epithelia via an apoptotic mechanism. Hepatology. 28, 906–913. Cicirello, H.G., Kehl, K.S., Addiss, D.G., Chusid, M.J., Glass, R.I., Davis, J.P. and Havens, P.L. 1997. Cryptosporidiosis in children during a massive waterborne outbreak in Milwaukee, Wisconsin: Clinical, laboratory and epidemiologic findings. Epidemiol. Infect. 119, 53–60. Clark, D.P. and Sears, C.L. 1996. The pathogenesis of cryptosporidiosis. Parasitol. Today. 12, 221–225. Clavel, A., Arnal, A.C., Sanchez, E.C., Cuesta, J., Letona, S., Amiguet, J.A., Castillo, F.J., Varea, M. and Gomez-Luz, R. 1996. Respiratory cryptosporidiosis: Case series and review of literature. Infection. 24, 341–346. Clayton, F., Heller, T. and Kotler, D.P. 1994. Variation in the enteric distribution of cryptosporidia in acquired immunodeficiency syndrome. Am. J. Clin. Pathol. 102, 420–425. Clemente, C.M., Caramori, C.A., Padula, P. and Rodrigues, M.A. 2000. Gastric cryptosporidiosis as a clue for the diagnosis of the acquired immunodeficiency syndrome. Arq. Gastroenterol. 37, 180–182. Collings, S. and Highton, J. 2004.Cryptosporidium reactive arthritis. N.Z. Med. J. 117, 1. Collins, J.M., Blanshard, C., Cramp, M., Gazzard, B. and Gleeson, JA. 1992. Case report: Pneumatosis intestinalis occurring in association with cryptosporidiosis and HIV infection. Clin. Radiol. 46, 410–411. Cordell, R.L. and Addiss, D.G. 1994. Cryptosporidiosis in child care settings: A review of the literature and recommendations for prevention and control. Pediatr Infect. Dis. J. 13,310–317. Review.



247

Cordell, R.L., Thor, P.M., Addiss, D.G., Theurer, J., Lichterman, R., Ziliak, S.R., Juranek, D.D. and Davis, J.P. 1997. Impact of a massive waterborne cryptosporidiosis outbreak on child care facilities in metropolitan Milwaukee, Wisconsin. Pediatr. Infect. Dis. J. 16, 639–644. Craun, G.F. and Calderon, R.L. 2006. Observational epidemiologic studies of endemic waterborne risks: Cohort, case-control, time-series, and ecologic studies. J. Water Health. 4 Suppl 2, 101–119. Cron, R.Q. and Sherry, D.D. 1995. Reiter’s syndrome associated with cryptosporidial gastroenteritis. J. Rheumatol. 22, 1962–1963. Current, W.L., Reese, N.C., Ernst, J.V., Bailey, W.S., Heyman, M.B. and Weinstein, W.M. 1983. Human cryptosporidiosis in immunocompetent and immunodeficient persons. N. Engl. J. Med. 308, 1252. Current, W.L. and Reese, N.C. 1986. A comparison of endogenous development of three isolates of Cryptosporidium in suckling mice. J. Protozool. 33, 98–108. Current, W.L. and Haynes, T.B. 1984. Complete development of Cryptosporidium in cell culture. Science. 224, 603–605. Deng, M., Lancto, C.A. and Abrahamsen, M.S. 2004. Cryptosporidium parvum regulation of human epithelial cell gene expression. Int. J. Parasitol. 34, 73–82. Dillingham, R.A., Lima, A.A. and Guerrant, R.L. 2002. Cryptosporidiosis: Epidemiology and impact. Microbes Infect. 4,1059–1066. Review. Dimicoli, S., Bensoussan, D., Latger-Cannard, V., Straczek, J., Antunes, L., Mainard, L., Dao, A., Barbe, F., Araujo, C., Clement, L., Feugier, P., Lecompte, T., Stoltz, J.F. and Bordigoni, P. 2003. Complete recovery from Cryptosporidium parvum infection with gastroenteritis and sclerosing cholangitis after successful bone marrow transplantation in two brothers with X-linked hyper-IgM syndrome. Bone Marrow Transplant. 32, 733–737. DuPont, H.L., Chappell, C.L., Sterling, C.R., Okhuysen, P.C., Rose, J.B. and Jakubowski. W. 1995. The infectivity of Cryptosporidium parvum in healthy volunteers. N. Engl. J. Med. 332,855–859. Ernest, J.A., Blagburn, B.L., Lindsay, D.S. and Current, W.L. 1986. Infection dynamics of Cryptosporidium parvum (Apicomplexa: Cryptosporidiidae) in neonatal mice (Mus musculus). J. Parasitol 72:796–798. Fayer, R. and Ungar, B.L.P. 1986. Cryptosporidium spp. and cryptosporidiosis. Microbiol. Rev. 50, 458. Forbes, A., Blanchard, C., Gazzard, B. 1993. Natural history of AIDS related sclerosing cholangitis: A study of 20 cases. Gut. 34,116–121. Franco, R.M. and Cordeiro, Nda. S. 1996. Giardiasis and cryptosporidiosis in day-care centers in the municipality Campinas SP. Rev. Soc. Bras. Med. Trop. 29, 585–591. Fripp, P.J., Bothma, M.T., Crewe-Brown, H.H. 1991. Four years of cryptosporidiosis at GaRankuwa Hospital. J. Infect. 23, 93–100. Frost, F.J., Calderon, R.L., Muller, T.B., Curry, M., Rodman, J.S., Moss, D.M. and de la Cruz, A.A. 1998. A two-year follow-up survey of antibody to Cryptosporidium in Jackson County, Oregon following an outbreak of waterborne disease. Epidemiol. Infect. 121, 213–217. Frost FJ, Muller TB, Calderon RL, Craun GF. 2004. Analysis of serological responses to Cryptosporidium antigen among NHANES III participants. Ann. Epidemiol. 14, 473–478. Gambhir, I.S., Jaiswal, J,P. and Nath, G. 2003. Significance of Cryptosporidium as an aetiology of acute infectious diarrhea in elderly Indians. Trop. Med. Int. Health. 8, 415–419. Garcia Velarde, E, Chavez Legaspi, M., Coello Ramirez, P., Gonzalez, J. and Aguilar Benavides, S. 1991. Cryptosporidium sp. in 300 children with and without diarrhea. Arch. Invest. Med. (Mex). 22, 329–332. Garone, M.A.B., Winston, B.J. and Lewis, J.H. 1986. Cryptosporidiosis of the stomach. Am. J. Gastroenterol. 81, 465–470. Garza, D.H., Fedorak, R.N. and Soave, R. 1986. Enterotoxin-like activity in cultured cryptosporidia: Role in diarrhea. Gastroenterology. 90, 1424. Gatei, W., Wamae, C.N., Mbae, C., Waruru, A., Mulinge, E., Waithera, T., Gatika, S.M., Kamwati, S.K., Revathi, G., Hart, C.A. 2006. Cryptosporidiosis: Prevalence, genotype analysis, and symptoms associated with infections in children in Kenya. Am. J. Trop. Med. Hyg. 75, 78–82. Gentile, G., Venditti, M., Micozzi, A., Caprioli, A., Donelli, G., Tirindelli, C., Meloni, G., Arcese, W. and Martino, P. 1991. Cryptosporidiosis in patients with hematologic malignancies. Rev. Infect. Dis. 13,842–846.


248


Goddard, E.A., Mouton, S.C., Westwood, A.T., Ireland, J.D. and Durra, G. 2000. Cryptosporidiosis of the gastrointestinal tract associated with sclerosing cholangitis in the absence of documented immunodeficiency: Cryptosporidium parvum and sclerosing cholangitis in an immunocompetent child. J. Pediatr. Gastroenterol. Nutr. 31, 317–320. Godwin, T.A. 1991. Cryptosporidiosis in the acquired immunodeficiency syndrome: A study of 15 autopsy cases. Hum. Patho. 22, 1215–1224. Goldstein, S.T., Juranek, D.D., Ravenholt, O., Hightower, A.W., Martin, D.G., Mesnik, J.L., Griffiths, S.D., Bryant, A.J., Reich, R.R. and Herwaldt, B.L. 1996. Cryptosporidiosis: An outbreak associated with drinking water despite state-of-the-art water treatment. Ann Intern Med. 124,459–68. Erratum in: Ann. Intern. Med. 1996 Jul 15;125(2):158. Goncalves, E.M., da Silva, A.J., Eduardo, M.B., Uemura, I.H., Moura, I.N., Castilho, V.L., and Corbett CE. 2006. Multilocus genotyping of Cryptosporidium hominis associated with diarrhea outbreak in a day care unit in Sao Paulo. Clinics. 61, 119–126. Epub 2006 Apr 25. Goodgame, R.W., Kimball, K., Ou, C.N., White, Jr., A.C., Genta, R.M., Lifschitz, C.H. and Chappell, C.L. 1995. Intestinal function and injury in acquired immunodeficiency syndrome-related cryptosporidiosis. Gastroenterology. 108, 1075–1082. Griffiths, J.K., Moore. R., Dooley, S., Keusch, G.T. and Tzipori, S. 1994. Cryptosporidium parvum infection of Caco-2 cell monolayers induces an apical monolayer defect, selectively increases transmonolayer permeability, and causes epithelial cell death. Infect. Immun. 62, 4506–4514. Guarino, A., Canani, R.B., Casola, A., Pozio, E., Russo, R., Bruzzese, E., Fontana, M. and Rubino, A. 1995. Human intestinal cryptosporidiosis: Secretory diarrhea and enterotoxic activity in Caco-2 cells. J. Infect. Dis. 171,976–983. Guarino, A., Canani, R.B., Pozio, E., Terracciano, L., Albano, F. and Mazzeo, M. 1994. Enterotoxic effect of stool supernatant of Cryptosporidium-infected calves on human jejunum. Gastroenterology. 106, 28–34. Guarino, A., Castaldo, A., Russo, S., Spagnuolo, M.I., Canani, R.B., Tarallo, L., DiBenedetto, L. and Rubino, A. 1997. Enteric cryptosporidiosis in pediatric HIV infection. J. Pediatr. Gastroenterol. Nutr. 25, 182–187. Guerrant, D.I., Moore, S.R., Lima, A.A., Patrick, P.D., Schorling, J.B. and Guerrant, R.L. 1999 .Association of early childhood diarrhea and cryptosporidiosis with impaired physical fitness and cognitive function four-seven years later in a poor urban community in northeast Brazil. Am. J. Trop. Med. Hyg. 61,707–713. Guerrant, R.L., Petri, W.A. and Weikel, C.S. 1990. Parasitic causes of diarrhea: An overview, in Pathophysiology of Secretory Diarrhea. E. Lebenthal, M. Duffey (Eds.). Raven Press, Boston. Guerrant, R.L.1997. An emerging highly infectious threat to our water supply. Infect Agents Dis. 3, 51–57. Hamedi, Y., Safa, O. and Haidari, M. Cryptosporidium infection in diarrheic children in southeastern Iran. 2005. Pediatr. Infect. Dis. J. 24, 86–88. Hart, C.A., Baxby, D. and Blundell, N. 1984. Gastro-enteritis due to Cryptosporidium: A prospective survey in a children's hospital. J Infect. 9,264–270. Hashmey, R., Smith, N.H., Cron, S., Graviss, E.A., Chappell, C.L. and White, A.C. Jr. 1997. Cryptosporidiosis in Houston, Texas. A report of 95 cases. Medicine (Baltimore). 76, 118–139. Hay, E.M., Winfield, J. and McKendrick, M.W. 1987. Reactive arthritis associated with cryptosporidium enteritis. Br. Med. J. (Clin. Res. Ed.). 295, 248. Heine, J., Pohlenz, J.F., Moon, H.W. and Woode, G.N. 1984. Enteric lesions and diarrhea in gnotobiotic calves monoinfected with Cryptosporidium species. J. Infect. Dis. 150:768–775. Hernandez, J., Lackner, A., Aye, P., Mukherjee, K., Tweardy, D.J., Mastrangelo, M.A., Joel, W., Griffiths, J., D’Souza, M., Dixit, S. and Robinson, P. 2006. Substance P is responsible for physiologic alterations such as increased chloride ion secretion and glucose-malabsorption in cryptosporidiosis. Infect. Immun. Dec. 11; Epub ahead of print. Hojlyng, N., Molbak, K., Jepsen, S. 1986. Cryptosporidium spp., a frequent cause of diarrhea in Liberian children. J. Clin. Microbiol. 23, 1109–1113. Houpt, E.R., Bushen, O.Y., Sam, N.E., Kohli, A., Asgharpour, A., Ng, C.T., Calfee, D.P. Guerrant, R.L., Maro, V., Ole-Nguyaine, S. and Shao, J.F. 2005. Short report: Asymptomatic Cryptosporidium hominis infection among human immunodeficiency virus-infected patients in Tanzania. Am. J. Trop. Med. Hyg. 73, 520–522. Hunter, P.R., Hughes, S., Woodhouse, S., Raj, N., Syed, Q., Chalmers, R.M., Verlander, N.Q. and Goodacre, J. 2004. Health sequelae of human cryptosporidiosis in immunocompetent patients. Clin. Infect. Dis. 39,504–510.



249

Hunter, P.R. and Nichols, G. 2002. Epidemiology and clinical features of Cryptosporidium infection in immunocompromised patients. Clin. Microbiol. Rev. 15,145–154. Iribarren, J.A., Castiella, A., Lobo, C., Lopez, P., von Wichmann, M.A., Arrizabalaga, J., Rodriguez-Arrondo, F.J. and Alzate, L.F. 1997. AIDS-associated cryptosporidiosis with antral narrowing. A new case. J. Clin. Gastroenterology 25, 693–694. Isaacs, D., Hunt, G.H., Phillips, A.D., Price, E.H., Raafat, F. and Walker-Smith, J.A. 1985. Cryptosporidiosis in immunocompetent children. J. Clin. Pathol. 38, 76. Javier Enriquez, F., Avila, C.R., Ignacio Santos, J., Tanaka-Kido, J., Vallejo, O. and Sterling, C.R. 1997. Cryptosporidium infections in Mexican children: Clinical, nutritional, enteropathogenic, and diagnostic evaluations. Am. J. Trop. Med. Hyg. 56, 254–257. Jokipii, L., Pohjola, S. and Jokipii, A.M. 1985. Cryptosporidiosis associated with traveling and giardiasis. Gastroenterology 89, 838–842. Jokipii, L., Pohjola, S., Valle, S.L.Jokipii, A.M. 1985. Frequency, multiplicity, and repertoire of intestinal protozoa in healthy homosexual men and in patients with gastrointestinal symptoms. Ann. Clin. Res. 17, 57–59. Jokipii, L. and Jokipii, A.M. 1986. Timing of symptoms and oocyst excretion in human cryptosporidiosis. N. Engl. J. Med. 3 15, 1643–1647. Kahn, W.A., Rogers, K.A., Karim, M.M., Ahmed, S., Hibberd, P.L., Calderwood, S.B. and Ryan, E.T. 2004. Cryptosporidiosis among Bangladeshi children with diarrhea: A prospective, matched, case-control study of clinical features, epidemiology and systemic antibody responses. Am. J. Trop. Med. Hyg. 71, 412–419. Kahn, D.G., Garfinkle, J.M., Klonoff, D.C., Pembrook, L.J and Morrow, D.J. 1987. Cryptosporidial and cytomegaloviral hepatitis and cholecystitis. Arch. Pathol. Lab. Med. 111,879–881. Kazlow, P.G., Shah, K., Benkov, K.J., Dische, R. and LeLeiko, N.S. 1986. Esophageal cryptosporidiosis in a child with acquired immune deficiency syndrome. Gastroenterology. 91, 1301–1303. Kelly, P., Feakins, R., Domizio, P., Murphy, J., Bevins, C., Wilson, J., McPhail, G., Poulsom, R. and Dhaliwal, W. 2004. Paneth cell granule depletion in the human small intestine under infective and nutritional stress. Clin. Exp. Immunol. 135,303–309. Kirkpatrick, B.D., Daniels, M.M., Jean, S.S., Pape, J.W., Karp, C., Littenberg, B., Fitzgerald, D.W., Lederman, H.M., Nataro, J.P. and Sears, C.L. 2002. Cryptosporidiosis stimulates an inflammatory intestinal response in malnourished Haitian children. J. Infect. Dis. 186, 94–101. Kirkpatrick, B.D., Noel, F., Rouzier, P.D., Powell, J.L., Pape, J.W., Bois, G., Alston, W.K., Larsson, C.J., Tenney, K., Ventrone, C., Powden, C., Sreenivasan, M., Sears, C.L. 2006. Childhood cryptosporidiosis is associated with a persistent systemic inflammatory response. Clin. Infect. Dis. 43:604–608. Epub 2006 Jul 26. Koch, K.L., Phillips, D.J., Aber, R.C. and Current, W.L. 1985. Cryptosporidiosis in hospital personnel. Ann. Intern. Med. 102, 593. Koch, K.L., Shankey, T.V., Weinstein, G.S., Dye, R.E., Abt, A.B., Current, W.L. and Eyster, M.E. 1983. Cryptosporidiosis in a patient with hemophilia, common variable hypogammaglobulinemia, and the acquired immunodeficiency syndrome. Ann. Intern. Med. 99, 337–340. Lee, J.K., Song, H.J. and Yu, J.R. 2005. Prevalence of diarrhea caused by Cryptosporidium parvum in nonHIV patients in Jeollanam-do, Korea. Kor. J. Parasitol. 43, 111–114. Lefkowitch, J.H., Krumholz, S., Feng-Chen, K.C., Griffin, P., Despommier, D. and Brasitus, T.A. 1984. Cryptosporidiosis of the human small intestine: A light and electron microscopic study. Hum. Pathol. Aug. 15(8):746–752. Lima, A.A., Silva, T.M., Gifoni, A.M., Barrett, L.J., McAuliffe, I.T., Bao, Y., Fox, J.W., Fedorko, D.P. and Guerrant, R.L. 1997. Mucosal injury and disruption of intestinal barrier function in HIV-infected individuals with and without diarrhea and cryptosporidiosis. Am. J. Gastroenterol. 92, 1861–1866. Lopez-Velez, R., Tarazona, A., Garcia Camacho, A., Gomez-Mampazo, E., Guerrero, A., Moneira, V. and Villanueva, R. 1995. Intestinal and extraintestinal cryptosporidiosis in AIDS patients. Eur. J. Clin. Microbiol. Infect. Dis. 14,677–681. Lorntz, B., Soares, A.M., Moore, S.R., Pinkerton, R., Gansneder, B., Bovbjerg, V.E., Guyatt, H., Lima, A.M. and Guerrant, R.L. 2006. Early childhood diarrhea predicts impaired school performance. Pediatr. Infect. Dis. J. 25, 513–520. Lumadue, J.A., Manabe, Y.C., Moore, R.D., Belitsos, P.C., Sears, C.L. and Clark, D.P. 1998. A clinicopathologic analysis of AIDS-related cryptosporidiosis. AIDS. 12, 2459–2466.


250


Macfarlane, D.E. and Horner-Bryce, J. 1987. Cryptosporidiosis in well-nourished and malnourished children. Acta. Paediatr. Scand. 76, 474–477. MacKenzie, W.R., Hoxie, N.J., Proctor, M.E., Gradus, M.S., Blair, K.A., Peterson, D.E., Kazmierczak, J.J., Addiss, D.G., Fox, K.R. and Rose, J.B.. 1994. A massive outbreak in Milwaukee of cryptosporidium infection transmitted through the public water supply. N. Engl. J. Med. 331,161–167. Erratum in: N. Engl. J. Med. Oct 13; 331(15):1035. MacKenzie, W.R., Schell, W.L., Blair, K.A., Addiss, D.G., Peterson, D.E., Hoxie, N.J., Kazmierczak, J.J., and Davis, J.P. 1995. Massive outbreak of waterborne cryptosporidium infection in Milwaukee, Wisconsin: Recurrence of illness and risk of secondary transmission. Clin. Infect. Dis. 21, 57–62. Manabe, Y.C., Clark, D.P., Moore, R.D., Lumadue, J.A., Dahlman, H.R., Belitsos, P.C., Chaisson, R.E. and Sears, C.L. 1998. Cryptosporidiosis in patients with AIDS: Correlates of disease and survival. Clin. Infect. Dis. 27, 536–542. Margulis, S.J., Honig, C.L., Soave, R., Govoni, A.F., Mouradian, J.A. and Jacobson, I.M. 1986. Biliary tract obstruction in the acquired immunodeficiency syndrome. Ann. Intern. Med. 105,207–210. Erratum in: Ann. Intern. Med. 1986, Oct.,105(4):634. Mathieu, E., Levy, D.A., Veverka, F., Parrish, M.K., Sarisky, J., Shapiro, N., Johnson, S., Handzel, T., Hightower, A., Xiao, L., Lee, Y.M., York, S., Arrowood, M., Lee, R. and Jones, J.L. 2004. Epidemiologic and environmental investigation of a recreational water outbreak caused by two genotypes of Cryptosporidium parvum in Ohio in 2000. Am. J. Trop. Med. Hyg. 71, 582–589. McCarty, M., Choudhri, A.H., Helbert, M. and Crofton, M.E. 1989. Radiological features of AIDS related cholangitis. Clin. Radiol. 40, 582–585. McCole, D.F., Eckmann, L., Laurent, F. and Kagnoff, M.F. 2000. Intestinal epithelial cell apoptosis following Cryptosporidium parvum infection. Infect. Immun. 68, 1710–1713. McMillan, A. and McNeillage, G.J.C.1984. Comparison of the sensitivity of microscopy and culture in the laboratory diagnosis of intestinal protozoal infection. J. Clin. Pathol. 37, 809–811. Meisel, J.L., Perera, D.R., Meligro, C. and Rubin, C.E. 1976. Overwhelming watery diarrhea associated with a Cryptosporidium in an immunosuppressed patient. Gastroenterology. 70, 1156–1160. Modigliani, R., Bories, C., Le Charpentier, Y., Salmeron, M., Messing, B., Galian, A., Rambaud, J.C., Lavergne, A., Cochand-Priollet, B. and Desportes, I. 1985. Diarrhoea and malabsorption in acquired immune deficiency syndrome: A study of four cases with special emphasis on opportunistic protozoan infestations. Gut. 26, 179–187. Molbak, K., Andersen, M., Aaby, P., Hojlyng, N., Jakobsen, M., Sodemann, M., and da Silva, A.P. 1997. Cryptosporidium infection in infancy as a cause of malnutrition: A community study from Guinea-Bissau, west Africa. Am. J. Clin. Nutr. 65,149–152. Molbak, K., Gottschau, A., Aaby, P., Hojlyng, N., Ingholt, L. and da Silva, A.P. 1994. Prolonged breast feeding, diarrhoeal disease, and survival of children in Guinea-Bissau. Br. Med. J. 308, 1403–1406. Molbak, K., Hojlyng, N., Gottschau, A., Sa, J.C., Ingholt, L., da Silva, A.P. and Aaby, P. 1993. Cryptosporidiosis in infancy and childhood mortality in Guinea Bissau, west Africa. Br. Med. J. 307, 417–420. Moon, A., Spivak, W. and Brandt, L.J. 1999. Cryptosporidium-induced gastric obstruction in a child with congenital HIV infection: Case report and review of the literature. J. Pediatr. Gastroenterol. Nutr. 28,108–111. Moon, H.W., Pohlenz, J.F.L., Woodmansee, D.B., Woode, G.N., Heine, J., Abel, S. and Jarvenin, J.A. 1985. Intestinal cryptosporidiosis: Pathogenesis and immunity. Microecol. Therap. 15, 103–120. Muller, T.B., Frost, F.J., Craun, G.F. and Calderon, R.L2001. Serological responses to Cryptosporidium infection. Infect. Immun. 69, 1974–1975. Naumova, E.N., Egorov, A.I., Morris, R.D. and Griffiths, J.K. 2003. The elderly and waterborne Cryptosporidium infection: Gastroenteritis hospitalizations before and during the 1993 Milwaukee outbreak. Emerg. Infect. Dis. 9, 418–425. Navin, T.R.1985. Cryptosporidiosis in humans: Review of recent epidemiologic studies. Eur. J. Epidemiol. 1, 77. Navin, T.R. and Hardy, A.M. 1987. Cryptosporidiosis in patients with AIDS. J. Infect. Dis. 155, 150. Navin, T.R. and Juranek, D.D. 1984. Cryptosporidiosis: Clinical, epidemiologic, and parasitologic review. Rev. Infect. Dis. 6, 313, Nchito, M., Kelly, P., Sianongo, S., Luo, N.P., Feldman, R., Farthing, M. and Baboo, KS. 1998. Cryptosporidiosis in urban Zambian children: An analysis of risk factors. Am. J. Trop. Med. Hyg. 59, 435–437.



251

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.


252


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.



253

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.


52268_C009.fm Page 255 Thursday, September 27, 2007 7:51 AM

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


256


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


Prophylaxis and Chemotherapy

257

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


258


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



259

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


260


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



261

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


262


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.


Prophylaxis and Chemotherapy b.

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


264

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



265

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


266


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



267

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


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


268


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



269


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


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

–


270


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


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



271


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


272



Age

Drug


Mature

Sulfameter Sulfamethazine Sulfamethizole Sulfamethoxazole Sulfamethoxypyridazine Sulfanilamide Sulfanilic acid Sulfanitran Sulfapyridine Sulfasalazine Sulfathiazole Sulfisomidine Sulfisoxizole


Mature


Mature

Azithromycin Clarithromycin Erythromycin Oleandomycin Spiramycin Azithromycin Spiramycin

Rat (immunosuppressed with dexamethasone) Rat (immunosuppressed with dexamethasone)

Mature

Arprinocid

Mature

Lasalocid

Monensin Salinomycin


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



273


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


274


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



275

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


276


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


1 day

Lasalocid Maduramicin Monensin Narasin Salinomycin Semduramicin Halofuginone Salinomycin Lasalocid Monensin


6 days

Garlic extract Diclazuril Toltrazuril


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.



277

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


278


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.



279

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.

References Allam, A.F. and Shehab, A.Y. 2002. Efficacy of azithromycin, praziquantel and mirazid in treatment of cryptosporidosis in school children. J. Egypt. Soc. Parasitol. 32, 969–978. Andreani, T., Modigliani, R., Charpentier, Y.L., Galian, A., Brouet, J.C., Liance, M., Lechance, J., Messing, B. and Vernisse, B. 1983. Acquired immunodeficiency with intestinal cryptosporidiosis: Possible transmission by Haitian whole blood. Lancet. 1, 1187–1191. Angus, K.W., 1990. Cryptosporidiosis in ruminants, in Cryptosporidiosis in Animals and Man, Dubey, J.P., Speer, C.A. and Fayer, R. (Eds). CRC Press, Boca Raton, FL, 83. Angus, K.W., Hutchison, G., Campbell, I. and Snodgrass, D.R. 1984. Prophylactic effects of anticoccidial drugs in experimental murine cryptosporidiosis. Vet. Rec. 114, 166–168. Anonymous. 1984. Update: Treatment of cryptosporidiosis in patients with acquired immunodeficiency syndrome (AIDS). MMWR CDC Surveill. Summ. 33, 117–119. Anonymous, 1999. Halocur Data Sheet. www.intervet.co.uk/Products_Public/Halocur/090_Product_Data sheet.asp. 2007. Armitage, K., Flanigan, T., Carey, J., Frank, I., MacGregor, R.R., Ross, P., Goodgame, R. and Turner, J. 1992. Treatment of cryptosporidiosis with paromomycin: A report of five cases. Arch. Intern. Med. 152, 2497–2499. Armson, A., Sargent, K., MacDonald, L.M., Finn, M.P., Thompson, R.C.A. and Reynoldson, J.A. 1999. A comparison of the effects of two dinitroanilines against Cryptosporidium parvum in vitro and in vivo in neonatal mice and rats. FEMS Immunol. Med. Microbiol. 26, 109–113. Arrowood, M.J., Jaynes, J.M. and Healey, M.C. 1991. In vitro activities of lytic peptides against sporozoites of Cryptosporidium parvum. Antimicrob. Agents Chemother. 35, 224–227. Arrowood, M.J., Xie, L.T. and Hurd, M.R. 1994. In vitro assays of maduramicin activity against Cryptosporidium parvum. J. Eukaryot. Microbiol. 41, 23S. Baishanbo, A., Gargala, G., Duclos, C., François, A., Rossignol, J.F., Ballet, J.J. and Favennec, L. 2006. Efficacy of nitazoxanide and paromomycin in biliary tract cryptosporidiosis in an immunosuppressed gerbil model. J. Antimicrob. Chemother. 57, 353–355. Barr, S.C., Jamrosz, G.F., Hornbuckle, W.E., Bowman, D.D. and Fayer, R. 1994. Use of paromomycin for treatment of cryptosporidiosis in a cat. J. Am. Vet. Med. Assoc. 205, 1742–1743. Barrier, M., Lacroix-Lamandé, S., Mancassola, R., Auray, G., Bernardet, N., Chaussé, A.-M., Uematsu, S., Akira, S. and Laurent, F. 2006. Oral and intraperitoneal administration of phosphorothioate oligodeoxynucleotides leads to control of Cryptosporidium parvum infection in neonatal mice. J. Infect. Dis. 193, 1400–1407. Bissuel, F., Cotte, L., Rabodonirina, R.P., Piens, M.A. and Trepo, C. 1994. Paromomycin: An effective treatment for cryptosporidial diarrhea in patients with AIDS. Clin. Infect. Dis. 18, 447–449. Blagburn, B.L., Drain, K.L., Land, T.M., Kinard, R.G., Moore, P.H., Lindsay, D.S., Patrick, D.A., Boykin, D.W. and Tidwell, R.R. 1998. Comparative efficacy evaluation of dicationic carbazole compounds, nitazoxanide and paromomycin against Cryptosporidium parvum infections in a neonatal mouse model. Antimicrob. Agents Chemother. 42, 2877–2882.


280


Blagburn, B.L. and Soave, R., 1997. Prophylaxis and chemotherapy: Human and Animal, in Cryptosporidium and cryptosporidiosis, 2nd Edition, Fayer, R. (Ed). CRC Press, Boca Raton, FL, 111–128. Blagburn, B.L., Sundermann, C.A., Lindsay, D.S., Hall, J.E. and Tidwell, R.R. 1991. Inhibition of Cryptosporidium parvum in neonatal Hsd:(ICR)BR Swiss mice by polyether ionophores and aromatic amidines. Antimicrob. Agents Chemother. 35, 1520–1523. Blanshard, C., Shanson, D.C. and Gazzard, B.G. 1997. Pilot studies of azithromycin, letrazuril and paromomycin in the treatment of cryptosporidiosis. Int. J.STD AIDS. 8, 124–129. Bouche, H., Housse, T.C., Dumont, J.L., Carnot, F., Menu, Y., Aveline, B., Belghiti, J., Boboc, B., Erlinger, S., Berthelot, P. and Pol, S. 1993. AIDS-related cholangitis: Diagnostic features and course in 15 patients. J. Hepatol. 17, 34–39. Brasseur, P., Lemeteil, D. and Ballet, J.J. 1991. Anti-cryptosporidial drug activity screened with an immunosuppressed rat model. J. Protozool. 38, 230S–231S. Brasseur, P., Lemeteil, D. and Ballet, J.J. 1993. Curative and preventive anticryptosporidium activities of sinefungin in an immunosuppressed adult rat model. Antimicrob. Agents Chemother. 37, 889–892. Bushen, O.Y., Davenport, J.A., Lima, A.B., Piscitelli, S.C., Uzgiris, A.J., Silva, T.M.J., Leite, R., Kosek, M., Dillingham, R.A., Girao, A., Lima, A.A.M. and Guerrant, R.L. 2004. Diarrhea and reduced levels of antiretroviral drugs: Improvement with glutamine or alanyl-glutamine in a randomized controlled trial in Northeast Brazil. Clin. Infect. Dis. 38, 1764–1770. Cacciò, S.M. and Pozio, E. 2006. Advances in the epidemiology, diagnosis and treatment of cryptosporidiosis. Expert Rev. Anti Infect. Ther. 4, 429–443. Cama, V.A., Marshall, M.M., Shubitz, L.F., Ortega, Y.R. and Sterling, C.R. 1994. Treatment of acute and chronic Cryptosporidium parvum infections in mice using clarithromycin and 14-OH clarithromycin. J. Eukaryot. Microbiol. 41, 25S. Carr, A., Marriott, D., Field, A., Vasak, E. and Cooper, D.A. 1998. Treatment of HIV-1-associated microsporidiosis and cryptosporidiosis with combination antiretroviral therapy. Lancet. 351, 256–261. Castro-Hermida, J.A., Gómez-Couso, H., Ares-Mazás, M.E., Gonzalez-Bedia, M.M., Castañeda-Cancio, N., Otero-Espinar, F.J. and Blanco-Mendez, J. 2004a. Anticryptosporidial activity of furan derivative G1 and its inclusion complex with beta-cyclodextrin. J. Pharm. Sci. 93, 197–206. Castro-Hermida, J.A., Pors, I., Otero-Espinar, F., Luzardo-Alvarez, A., Ares-Mazás, E. and Chartier, C. 2004b. Efficacy of α-cyclodextrin against experimental cryptosporidiosis in neonatal goats. Vet. Parasitol. 120, 35–41. Castro-Hermida, J.A., Quílez-Cinca, J., López-Bernad, F., Sánchez-Acedo, C., Freire-Santos, F. and AresMazás, E. 2001. Treatment with β-cyclodextrin of natural Cryptosporidium parvum infections in lambs under field conditions. Int. J. Parasitol. 31, 1134–1137. Cello, J.P., Grendell, J.H., Basuk, P., Simon, D., Weiss, L., Wittner, M., Rood, R.P., Wilcox, M., Forsmark, C.E., Read, A.E., Satow, J.A., Weikel, C.S. and Beaumont, C. 1991. Effect of octreotide on refractory AIDS-associated diarrhea: a prospective, multicenter clinical trial. Ann. Intern. Med. 115, 705–710. Chandrasekar, P.H. 1987. “Cure” of chronic cryptosporidiosis during treatment with azidothymidine in a patient with acquired immunodeficiency syndrome. Am. J. Med. 83, 187. Clezy, K., Gold, J., Blaze, J. and Jones, P. 1991. Paromomycin for the treatment of cryptosporidial diarrhea in AIDS patients. AIDS 5, 1146–1147. Clotet, B., Sirera, G., Cofran, F., Monterola, J.M., Tortosa, F. and Fox, M. 1989. Efficacy of the somatostatin analogue (SMS-201-995), Sandostatin, for cryptosporidial diarrhea in patients with AIDS. AIDS 3, 857–858. Collier, A.C., Miller, R. and Meyers, J. 1984. Cryptosporidiosis after marrow transplantation: Person-to-person transmission and treatment with spiramycin. Ann. Intern. Med. 101, 205–206. Connolly, G.M., Dryden, M.S., Shanson, D.C. and Gazzard, B.G. 1988. Cryptosporidial diarrhea in AIDS and its treatment. Gut. 29, 593–597. Connolly, G.M., Youle, M. and Gazzard, B.G. 1990. Diclazuril in the treatment of severe cryptosporidial diarrhea in AIDS patients. AIDS 4, 700–701. Cranfield, M.R. and Graczyk, T.K. 1994. Experimental infection of elaphid snakes with Cryptosporidium serpentis (Apicomplexa: Cryptosporidiidae). J. Parasitol. 80, 823–826. Current, W.L. and Garcia, L.S. 1991. Cryptosporidiosis. Clin. Microbiol. Rev. 4, 325–358. Danziger, L.H., Kanyok, T.P. and Novak, R.M. 1993. Treatment of cryptosporidial diarrhea in an AIDS patient with paromomycin. Ann. Pharmacother. 27, 1460–1462.



281

Dionisio, D., Orsi, A., Sterrantino, G., Meli, M., Lollo, S.D., Manneschi, L.I., Trotta, M., Pozzi, M., Sani, L. and Leoncini, F. 1998. Chronic cryptosporidiosis in patients with AIDS: Stable remission and possible eradication after long term, low dose azithromycin. J. Clin. Pathol. 51, 138–142. Doumbo, O., Rossignol, J.F., Prichard, E., Traore, H., Dembele, M., Diakite, M., Traore, F. and Dialio, A.D. 1997. Nitazoxanide in the treatment of cryptosporidial diarrhea and other intestinal parasitic infection associated with Acquired Immunodeficiency Syndrome in tropical Africa. Am. J. Trop. Med. Hyg. 56, 637–639. Fafard, J. and Lalonde, R. 1990. Long-standing symptomatic cryptosporidiosis in a normal man: Clinical response to spiramycin. J. Clin. Gastroenterol. 12, 190–191. Fanning, M., Monte, M., Sutherland, L.R., Broadhead, M., Murphy, G.F. and Harris, A.G. 1991. Pilot study of Sandostatin (octreotide) therapy of refractory HIV-associated diarrhea. Dig. Dis. Sci. 36, 476–480. Farr, B.M., 2000. Rifamycins, in Principles and Practice of Infectious Diseases, 5th Edition, Mandell, G.L., Bennett, J.E. and Dolin, R. (Eds). Churchill Livingstone, Philadelphia, PA, 1534. Favennec, L., Egraz-Bernard, M., Comby, E., Lemeteil, D., Ballet, J.J. and Brasseur, P. 1994. Immunofluorescence detection of Cryptosporidium parvum in Caco-2 cells: A new screening method for anticryptosporidial agents. J. Eukaryot. Microbiol. 41, 39S. Fayer, R. 1992. Activity of sulfadimethoxine against cryptosporidiosis in dairy calves. J. Parasitol. 78, 534–537. Fayer, R. and Ellis, W. 1993a. Glycoside antibiotics alone and combined with tetracyclines for prophylaxis of experimental cryptosporidiosis in neonatal BALB/c mice. J. Parasitol. 79, 553–558. Fayer, R. and Ellis, W. 1993b. Paromomycin is effective as prophylaxis for cryptosporidiosis in dairy calves. J. Parasitol. 79, 771–774. Fayer, R. and Ellis, W. 1994. Qinghaosu (Artemisinin) and derivatives fail to protect neonatal BALB/c mice against Cryptosporidium parvum (Cp) infection. J. Eukaryot. Microbiol. 41, 41S. Fayer, R., Phillips, L., Anderson, B.C. and Bush, M. 1991. Chronic cryptosporidiosis in a Bactrian camal (Camelus bactrianus). J. Zoo Wildl. Med. 22, 228–232. Fayer, R. and Ungar, B.L. 1986. Cryptosporidium spp. and cryptosporidiosis. Microbiol. Rev. 50, 458–483. Ferre, I., Benito-Peña, A., García, U., Osoro, K. and Ortega-Mora, L.M. 2005. Effect of different decoquinate treatments of cryptosporidiosis in naturally infected cashmere goat kids. Vet. Rec. 157, 261–262. Fichtenbaum, C.J., Ritchie, D.J. and Powderly, W.G. 1993. Use of paromomycin for treatment of cryptosporidiosis in patients with AIDS. Clin. Infect. Dis. 16, 298–300. Fichtenbaum, C.J., Zackin, R., Feinberg, J., Benson, C. and Griffiths, J.K. 2000. Rifabutin but not clarithromycin prevents cryptosporidiosis in persons with advanced HIV infection. AIDS 14, 2889–2893. Fischer, O. 1983. Attempted therapy and prophylaxis of cryptosporidiosis in calves by administration of sulphadimidine. Acta Vet. Brno. 52, 183. Foudraine, N.A., Weverling, G.J., Gool, T.V., Roos, M.T.L., Wolf, F.D., Koopmans, P.P., Broek, P.J.V.D., Meenhorst, P.L., Leeuwen, R.V., Lange, J.M.A. and Reiss, P. 1998. Improvement of chronic diarrhoea in patients with advanced HIV-I infection during potent antiretroviral therapy. AIDS 12, 35–41. Fox, L.M. and Saravolatz, L.D. 2005. Nitazoxanide: A new thiazolide antiparasitic agent. Clin. Infect. Dis. 40, 1173–1180. Freid, E., 2005. FDA approves first-ever treatment for adults, teens with diarrhea caused by Cryptosporidium infection. www.alinia.com/index.php/media_center.html. Accessed 2006. Gamra, M.M.M.A. and Hosseiny, L.M.E. 2003. Comparative study of the prophylactic and therapeutic effects of paromomycin, recombinant IL-12 alone or in combination against Cryptosporidium parvum infection in immunosuppressed mice. J. Egypt. Soc. Parasitol. 33, 109–122. Gargala, G., Baishanbo, A., Favennec, L., François, A., Ballet, J.J. and Rossignol, J.F. 2005. Inhibitory activities of epidermal growth factor receptor tyrosine kinase-targeted dihydroxyisoflavone and trihydroxydeoxybenzoin derivatives on Sarcocystis neurona, Neospora caninum, and Cryptosporidium parvum development. Antimicrob. Agents Chemother. 49, 4628–4634. Gathe, J., Piot, D., Hawkins, K., Bernal, A., Clemmons, J. and Stool, E. 1990. Treatment of gastrointestinal cryptosporidiosis with paromomycin. International Conference on AIDS, Abstract 2121, San Francisco, CA. Georgiev, V. 1993. Opportunistic infections: Treatment and developmental therapeutics of cryptosporidiosis and isosporiasis. Drug Dev. Res. 28, 445–459.


282


Giacometti, A., Cirioni, O. and Scalise, G. 1996. In-vitro activity of macrolides alone and in combination with artemisin, atovaquone, dapsone, minocycline or pyrimethamine against Cryptosporidium parvum. J. Antimicrob. Chemother. 38, 399–408. Girard, P., Goldschmidt, E., Vittecoq, D., Massip, P., Gastiaburu, J., Meyohas, M.C., Couland, J.P. and Schally, A.V. 1992. Vapreotide, a somatostatin analogue, in cryptosporidiosis and other AIDS-related diarrheal diseases. AIDS 6, 715–718. Glisson, J.R., Brown, T.P., Brugh, M., Page, R.K., Kleven, S.H. and Davis, R.B. 1984. Sinusitis in turkeys associated with respiratory cryptosporidiosis. Avian Dis. 28, 783–790. Gobel, E. 1987a. Diagnose und therapie der akuten Kryptosporidiose beim kalb. Tieraerztl. Umsch. 42, 863–869. Gobel, E. 1987b. Possibilities of therapy of cryptosporidiosis in calves in problematic farms. Zentralbl. Bakteriol. Mikrobiol. Hyg [A]. 265, 489. Gobel, E. and Bretschneider, M. 1985. Mikromorphologische untersuchungen zur wirksamkeit von lasalocid auf die entwicklungsstadien von Cryptosporidium. Kongresses der Deuteschen Veterinarmedizinischen Gesellschaft, 278–282, Bad Nauheim. 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., Helmer, P., Fayer, R. and Bostwick, E.F. 1998. Therapeutic efficacy of hyperimmune bovine colostrum treatment against clinical and subclinical Cryptosporidium serpentis infections in captive snakes. Vet. Parasitol. 74, 123–132. Greenberg, R.E., Mir, R., Bank, S. and Siegal, F.P. 1989. Resolution of intestinal cryptosporidiosis after treatment of AIDS with AZT. Gastroenterology 97, 1327–1330. Gross, T.L., Wheat, J., Bartlett, M. and O’Connor, K.W. 1986. AIDS and multiple system involvement with Cryptosporidium. Am. J. Gastroenterol. 81, 456–458. Guillem, S., Gomez, M., Romeu, J., Raventos, A., Fernandez, A., Condom, M.J. and Clotet, B. 1992. Letrazuril for treatment of severe cryptosporidial diarrhea in AIDS. 8th International Conference on AIDS/III STD World Congress, Abstract PoB3257, Amsterdam. Guitard, J., Menotti, J., Desveaux, A., Alimardani, P., Porcher, R., Derouin, F. and Kapel, N. 2006. Experimental study on the effects of probiotics on Cryptosporidium parvum infection in neonatal rats. Parasitol. Res. 99, 522–527. Gutteridge, W.E. 1991. 566C80, an antimalarial hydroxynaphthoquinone with broad spectrum experimental activity against opportunistic parasitic infections of AIDS patients. J. Protozool. 38, 141S–143S. Hamour, A.A., Bonington, A., Hawthorne, B. and Wilkins, E.G.L. 1993. Successful treatment of AIDS-related cryptosporidial sclerosing cholangitis. AIDS 7, 1449–1451. Harp, J.A. 1999. Oral dosing of neonatal mice with sucrose reduces infection with Cryptosporidium parvum. J. Parasitol. 85, 952–955. Harris, M., Deutsch, G., Maclean, J.D. and Tsoukas, C.M. 1994. A phase I study of letrazuril in AIDS-related cryptosporidiosis. AIDS 8, 1109–1113. Healey, M.C., Yang, S., Rasmussen, K.R., Jackson, M.K. and Du, C. 1995. Therapeutic efficacy of paromomycin in immunosuppressed adult mice infected with Cryptosporidium parvum. J. Parasitol. 81, 114–116. Hewitt, R.G., Yiannoutsos, C.T., Higgs, E.S., Carey, J.T., Geiseler, P.J., Soave, R., Rosenberg, R., Vazquez, G.J., Wheat, L.J., Fass, R.J., Antoninievic, Z., Walawander, A.L., Flanigan, T.P. and Bender, J.F. 2000. Paromomycin: No more effective than placebo for treatment of cryptosporidiosis in patients with advanced human immunodeficiency virus infection. Clin. Infect. Dis. 31, 1084–1092. Hijjawi, N.S., Meloni, B.P., Ng’anzo, M., Ryan, U.M., Olson, M.E., Cox, P.T., Monis, P.T. and Thompson, R.C.A. 2004. Complete development of Cryptosporidium parvum in host cell-free culture. Int. J. Parasitol. 34, 769–777. Hinnant, K., Swartz, A., Rotterdam, H., Bell, E. and Tappen, M. 1989. Cytomegaloviral and cryptosporidial cholecystitis in two patients with AIDS. Am. J. Surg. Pathol. 13, 57–60. Hoerr, F.J., Current, W.L. and Haynes, T.B. 1986. Fatal cryptosporidiosis in quail. Avian Dis. 30, 421–425. Hoffmann, C., 2006. Cryptosporidiosis, in HIV Medicine 2006, 14th Edition, 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 Edition, Hoffmann, C., Rockstroh, J. and Kamps, B.S. (Eds). Flying Publisher, 825.



283

Hoffmann, C. and Mulcahy, F., 2006b. Nucleoside analogs (“nukes,” NRTIs), in HIV Medicine 2006, 14th Edition, 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 Edition, Hoffmann, C., Rockstroh, J. and Kamps, B.S. (Eds). Flying Publisher, 825. Holmberg, S.D., Moorman, A.C., Bargen, J.C.V., Palella, F.J., Loveless, M.O., Ward, D.J. and Navin, T.R. 1998. Possible effectiveness of clarithromycin and rifabutin for cryptosporidiosis chemoprophylaxis in HIV disease. JAMA 279, 384–386. Hommer, V., Eichholz, J. and Petry, F. 2003. Effect of antiretroviral protease inhibitors alone, and in combination with paromomycin, on the excystation, invasion and in vitro development of Cryptosporidium parvum. J. Antimicrob. Chemother. 52, 359–364. Huang, K. and Yang, S. 2002. Inhibitory effect of selenium on Cryptosporidium parvum infection in vitro and in vivo. Biol. Trace Elem. Res. 90, 261–272. Hunt, E., Fu, Q., Armstrong, M.U., Rennix, D.K., Webster, D.W., Galanko, J.A., Chen, W., Weaver, E.M., Argenzio, R.A. and Rhoads, J.M. 2002. Oral bovine serum concentrate improves cryptosporidial enteritis in calves. Pediatr. Res. 51, 370–376. Ives, N.J., Gazzard, B.G. and Easterbrook, P.J. 2001. The changing pattern of AIDS-defining illnesses with the introduction of Highly Active Antiretroviral Therapy (HAART) in a London clinic. J. Infect. 42, 134–139. Joachim, A., Krull, T., Schwarzkopf, J. and Daugschies, A. 2003. Prevalence and control of bovine cryptosporidiosis in German dairy herds. Vet. Parasitol. 112, 277–288. Jordan, W.C. 1996. Clarithromycin prophylaxis against cryptosporidium enteritis in patients with AIDS. J. Natl. Med. Assoc. 88, 425–427. Kadappu, K.K., Nagaraja, M.V., Rao, P.V. and Shastry, B.A. 2002. Azithromycin as treatment for cryptosporidiosis in human immunodeficiency virus disease. J. Postgrad. Med. 48, 179–181. Kamps, B.S. and Hoffman, C., 2006. Drugs, in HIV Medicine 2006, 14th Edition, Hoffman, C., Rockstroh, J. and Kamps, B.S. (Eds). Flying Publisher, 825. Kern, P., Toy, J. and Dietrich, M. 1985. Preliminary clinical observation with recombinant interleukin-2 in patients with AIDS or LAS. Blut. 50, 1–6. Kim, C.W. 1987. Chemotherapeutic effect of arprinocid in experimental cryptosporidiosis. J. Parasitol. 73, 663–666. Kimata, I., Shigehiko, U. and Iseki, M. 1991. Chemotherapeutic effect of azithromycin and lasalocid on Cryptosporidium infection in mice. J. Protozool. 38, 232S–233S. Kobayashi, C., Yokoyama, H., Nguyen, S.V., Kodama, Y., Kimata, T. and Izeki, M. 2004. Effect of egg yolk antibody on experimental Cryptosporidium parvum infection in scid mice. Vaccine. 23, 232–235. Kuhls, T.L., Mosier, D.A., Abrams, V.L., Crawford, D.L. and Greenfield, R.A. 1994. Inability of interferongamma and aminoguanidine to alter Cryptosporidium parvum infection in mice with severe immunodeficiency. J. Parasitol. 80, 480–485. Lallemond, M., Villeneuve, A., Belda, J. and Dubreuil, P. 2006. Field study of the efficacy of halofuginone and decoquinate in the treatment of cryptosporidiosis in veal calves. Vet. Rec. 159, 672–677. Leitch, G.J. and He, Q. 1994a. Arginine-derived nitric-oxide reduces fecal oocyst shedding in nude mice infected with Cryptosporidium parvum. Infect. Immun. 62, 5173–5176. Leitch, G.J. and He, Q. 1994b. Putative anticryptosporidial agents tested in an immunodeficient mouse model. Antimicrob. Agents Chemother. 38, 865–867. Lemeteil, D., Roussel, R., Favennec, L., Ballet, J.J. and Brasseur, P. 1993. Assessment of candidate anticryptosporidial agents in an immunosuppressed rat model. J. Infect. Dis. 167, 766–768. Lindsay, D.S., Blagburn, B.L., Sundermann, C.A. and Ernest, J.A. 1987. Chemoprophylaxis of cryptosporidiosis in chickens using halofuginone, salinomycin, lasalocid, or monensin. Am. J. Vet. Res. 48, 354–355. Loeb, M., Walach, C., Phillips, J., Fong, I., Salit, I., Rachlis, A. and Walmsley, S. 1995. Treatment with letrazuril of refractory cryptosporidial diarrhea complicating AIDS. J. Acquir. Immune Defic. Syndr. Hum. Retrovirol. 10, 48–53. Maggi, P., Larocca, A.M.V., Quarto, M., Serio, G., Brandonisio, O., Angarano, G. and Pastore, G. 2000. Effect of antiretroviral therapy on cryptosporidiosis and microsporidiosis in patients infected with Human Immunodeficiency Virus Type I. Eur. J. Clin. Microbiol. Infect. Dis. 19, 213–217.


284


Mancassola, R., Reperant, J.M., Naciri, M. and Chartier, C. 1995. Chemoprophylaxis of Cryptosporidium parvum infection with paromomycin in kids and immunological study. Antimicrob. Agents Chemother. 39, 75–78. Mancassola, R., Richard, A. and Naciri, M. 1997. Evaluation of decoquinate to treat experimental cryptosporidiosis in kids. Vet. Parasitol. 69, 31–37. Mannheimer, S.B. and Soave, R. 1994. Protozoal infections in patients with AIDS. Infect. Dis. Clin. North Am. 8, 483–498. Marshall, R.J. and Flanigan, T.P. 1992. Paromomycin inhibits Cryptosporidium infection of a human enterocyte cell line. J. Infect. Dis. 165, 772–774. Mason, R.W. and Hartley, W.J. 1980. Respiratory cryptosporidiosis in a peacock chick. Avian Dis. 24, 771–776. McDonald, S.A.C., O’Grady, J.E., Bajaj-Elliott, M., Notley, C.A., Alexander, J., Brombacher, F. and McDonald, V. 2004. Protection against the early acute phase of Cryptosporidium parvum infection conferred by interleukin-4-induced expression of T helper 1 cytokines. J. Infect. Dis. 190, 1019–1025. McDonald, V., Stables, R., Warhurst, D.C., Barer, M.R., Blewett, D.A., Chapman, D.H., Connolly, G.M., Chiodini, P.L. and McAdam, K.P.W.J. 1990. In vitro cultivation of Cryptosporidium parvum and screening for anticryptosporidial drugs. Antimicrob. Agents Chemother. 34, 1498–1500. Mead, J.R., You, X., Pharr, J.E., Belenkaya, Y., Arrowood, M.J., Fallon, M.T. and Schinazi, R.F. 1995. Evaluation of maduramicin and alborixin in a SCID mouse model of chronic cryptosporidiosis. Antimicrob. Agents Chemother. 39, 854–850. Meamar, A.-R., Rezaian, M., Rezaie, S., Mohraz, M., Kia, E.B., Houpt, E.R. and Solaymani-Mohammadi, S. 2006. Cryptosporidium parvum bovine genotype oocysts in the respiratory samples of an AIDS patient: Efficacy of treatment with a combination of azithromycin and paromomycin. Parasitol Res. 98, 593–595. Mele, R., Morales, M.A.G., Tosini, F. and Pozio, E. 2003. Indinavir reduces Cryptosporidium parvum infection in both in vitro and in vivo models. Int. J. Parasitol. 33, 757–764. Menichetti, F., Moretti, M.V., Marroni, M., Papili, R. and DiCandilo, F. 1991. Diclazuril for cyptosporidiosis in AIDS. Am. J. Med. 90, 271–272. Miao, Y.M., Awad-El-Kariem, F.M., Gibbons, C.L. and Gazzard, B.G. 1999. Cryptosporidiosis: Eradication or suppression with combination antiretroviral therapy? AIDS 13, 734–735. Miller, T.L., Winter, H.S., Luginbuhl, L.M., Orav, E.J. and McIntosh, K. 1992. Pancreatitis in pediatric human immunodeficiency virus infection. J. Pediatr. 120, 223–227. Moling, O., Avi, A., Rimenti, G. and Mian, P. 2005. Glutamine supplementation for patients with severe cryptosporidiosis. Clin. Infect. Dis. 40, 773–774. Moon, H.W., Schwartz, A., Welch, M.J., McCann, P.P. and Runnels, P.L. 1982a. Experimental fecal transmission of human cryptosporidia to pigs, and attempted treatment with an ornithine decarboxylase inhibitor. Vet. Pathol. 19, 700–707. Moon, H.W., Woode, G.N. and Ahrens, F.A. 1982b. Attempted chemoprophylaxis of cryptosporidiosis in calves. Vet. Rec. 110, 181. Moore, D., Waters, W.R., Wannemuehler, M.J. and Harp, J.A. 2001. Treatment with agmatine inhibits Cryptosporidium parvum infection in infant mice. J. Parasitol. 87, 211–213. Moore, D.A., Atwill, E.R., Kirk, J.H., Brahmbhatt, D., Alonso, L.H., Hou, L., Singer, M.D. and Miller, T.D. 2003. Prophylactic use of decoquinate for infections with Cryptosporidium parvum in experimentally challenged neonatal calves. J. Am. Vet. Med. Assoc. 223, 839–845. Moore, R.D. and Chaisson, R.E. 1999. Natural history of HIV infection in the era of combination antiretroviral therapy. AIDS 13, 1933–1942. Moroni, M., Esposito, R., Cernuschi, M., Franzetti, F., Carosi, G.P. and Fiori, G.P. 1993. Treatment of AIDSrelated refractory diarrhea with octreotide. Digestion. 54, 30–32. Morrison, D.A., Bornstein, S., Thebo, P., Wernery, U., Kinne, J. and Mattson, J.G. 2004. The current status of the small subunit rRNA phylogeny of the coccidian (Sporozoa). Int. J. Parasitol. 34, 501–514. Moskovitz, B.L., Stanton, T.L. and Kusmierek, J.J.E. 1988. Spiramycin therapy for cryptosporidial diarrhea in immunocompromised patients. J. Antimicrob. Chemother. 22, 189–191. Murdoch, D.A., Bloss, D.E. and Glover, S.C. 1993. Successful treatment of cryptosporidiosis in an AIDS patient with letrazuril. AIDS. 7, 1279–1280. Nachbaur, D., Kropshofer, G., Feichtinger, H., Allerberger, F. and Niederwieser, D. 1997. Cryptosporidiosis after CD34-selected autologous peripheral blood stem cell transplantation (PBSCT). Treatment with paromomycin, azithromycin and recombinant human interleukin-2. Bone Marrow Trans. 19, 1261–1263.



285

Naciri, M., Mancassola, R., Yvore, P. and Peeters, J.E. 1993. The effect of halofuginone lactate on experimental C. parvum infections in calves. Vet. Parasitol. 45, 199–207. Naciri, M. and Yvore, P. 1989. Efficite du lactate d’halofuginone dans le traitment de le cryptosporidiose chez l’agneau. Rec. Med. Vet. Ec. Alfort. 165, 823. Naciri, M., Yvore, P. and Levieux, D. 1984. Cryptosporidiose experimentale du chevreau. Influence de la prise du colostrum. Essais de traitments. Colloque International, Abstract 465, Niort. Nagy, B., Bozso, M., Palfi, V., Nagy, G. and Sahiby, M.A. 1984. Studies on cryptosporidial infection of goat kids. Colloque International, Abstract 443, Niort. O’Donoghue, P.J. 1995. Cryptosporidium and cryptosporidiosis in man and animals. Int. J. Parasitol. 25, 139–195. Palella, F.J., Delaney, K.M., Moorman, A.C., Loveless, M.O., Fuhrer, J., Satten, G.A., Aschman, D.J. and Holmberg, S.D. 1998. Declining morbidity and mortality among patients with advanced human immunodeficiency virus infection. N. Engl. J. Med. 338, 853–860. Pasquali, P., Fayer, R., Zarlenga, D., Canals, A., Marez, T.d., Munoz, M.T.G., Almeria, S. and Gasbarre, L.C. 2006. Recombinant bovine interleukin-12 stimulates a gut immune response but does not provide resistance to Cryptosporidium parvum infection in neonatal calves. Vet. Parasitol. 135, 259–268. Peeters, J.E., Villacorta, I., Naciri, M. and Vanopdenbosch, E. 1993. Specific serum and local antibody responses against Cryptosporidium parvum during medication of calves with halofuginone lactate. Infect. Immun. 61, 4440–4445. Petersen, C. 1992. Cryptosporidiosis in patients infected with the human immunodeficiency virus. Clin. Infect. Dis. 815, 903–909. Pickerd, N. and Tuthill, D. 2004. Resolution of cryptosporidiosis with probiotic treatment. Postgrad. Med. J. 80, 112–113. Portnoy, D., Whiteside, M.E., Buckley, E. and Macleod, C.L. 1984. Treatment of intestinal cryptosporidiosis with spiramycin. Ann. Intern. Med. 101, 202–204. Pozio, E. and Morales, M.A.G. 2005. The impact of HIV-protease inhibitors on opportunistic parasites. Trends Parasitol. 21, 58–63. Rasmussen, K.R., Arrowood, M.J. and Healey, M.C. 1992. Effectiveness of dehydroepiandrosterone in reduction of cryptosporidial activity in immunosuppressed rats. Antimicrob. Agents Chemother. 36, 220–222. Rasmussen, K.R., Healey, M.C., Cheng, L. and Yang, S. 1995. Effects of dehydroepiandrosterone in immunosuppressed adult mice infected with Cryptosporidium parvum. J. Parasitol. 81, 429–433. Rasmussen, K.R. and Healy, M.C. 1992. Dehydroepiandrosterone-induced reduction of Cryptosporidium parvum infection in aged Syrian golden hamsters. J. Parasitol. 78, 554–557. Rehg, J.E. 1991a. Anticryptosporidial activity is associated with specific sulfonamides in immunosuppressed rats. J. Parasitol. 77, 238–240. Rehg, J.E. 1991b. Anti-cryptosporidial activity of macrolides in immunosuppressed rats. J. Protozool. 38, 228S–230S. Rehg, J.E. 1991c. Activity of azithromycin against cryptosporidia in immunosuppressed rats. J. Infect. Dis. 163, 1293–1296. Rehg, J.E. 1993. Anticryptosporidial activity of lasalocid and other ionophorous antibiotics in immunosuppressed rats. J. Infect. Dis. 168, 1566–1569. Rehg, J.E. 1994. A comparison of anticryptosporidial activity of paromomycin with that of other aminoglycosides and azithromycin in immunosuppressed rats. J. Infect. Dis. 170, 934–938. Rehg, J.E. 1995. The activity of halofuginone in immunosuppressed rats infected with Cryptosporidium parvum. J. Antimicrob. Chemother. 35, 391–397. Rehg, J.E. 1996. Effect of diethyldithiocarbamate on Cryptosporidium parvum infections in immunosuppressed rats. J. Parasitol. 82, 158–162. Rehg, J.E. and Hancock, M.L. 1990. Effectiveness of arprinocid in the reduction of cryptosporidial activity in immunosuppressed rats. Am. J. Vet. Res. 51, 1668–1670. Rehg, J.E., Hancock, M.L. and Woodmansee, D.B. 1988. Anticryptosporidial activity of sulfadimethoxine. Antimicrob. Agents Chemother. 32, 1907–1908. Riggs, M.W., Schaefer, D.A., Kapil, S.J., Barley-Maloney, L. and Perryman, L.E. 2002. Efficacy of monoclonal antibodies against defined antigens for passive immunotherapy of chronic gastrointestinal cryptosporidiosis. Antimicrob. Agents Chemother. 46, 275–282.


286


Ritchie, D.J. and Becker, E.S. 1994. Update on the management of intestinal cryptosporidiosis in AIDS. Ann. Pharmacother. 28, 767–778. Rohlman, V.C., Kuhls, T.L., Mosier, D.A., Crawford, D.L., Hawkins, D.R., Abrams, V.L. and Greenfield, R.A. 1993. Therapy with atovaquone for Cryptosporidium parvum infection in neonatal severe combined immunodeficiency mice. J. Infect. Dis. 168, 258–260. Rolston, K.V.I., Fainstein, V. and Bodey, G.P. 1989. Intestinal cryptosporidiosis treated with eflornithine: A prospective study among patients with AIDS. JAIDS J. Acquir. Immune Defic. Syndr. 2, 426–430. Romeu, J., Miro, J.M., Sirera, G., Mallolas, J., Arnal, J., Valls, M.E., Tortosa, F., Clotet, B. and Foz, M. 1991. Efficacy of octreotide in the management of chronic diarrhea in AIDS. AIDS 5, 1495–1499. Rossignol, J.-F.A., Ayoub, A. and Ayers, M.S. 2001. Treatment of diarrhea caused by Cryptosporidium parvum: A prospective randomized, double-blind, placebo-controlled study of nitazoxanide. J. Infect. Dis. 184, 103–106. Rossignol, J.-F.A., Hidalgo, H., Feregrino, M., Higuera, F., Gomez, W.H., Romero, J.L., Padierna, J., Geyne, A. and Ayers, M.S. 1998. A double-“blind” placebo-controlled study of nitazoxanide in the treatment of cryptosporidial diarrhoea in AIDS patients in Mexico. Trans. Roy. Soc. Trop. Med. Hyg. 92, 663–666. Rossignol, J.F. and Maisonneuve, H. 1984. Nitazoxanide in the treatment of Taenia saginata and Hymenolepis nana infections. Am. J. Trop. Med. Hyg. 33, 511–512. Rossignol, J.F., Maisonneuve, H. and Cho, Y.W. 1984. Nitroimidazoles in the treatment of trichomoniasis, giardiasis and amoebiasis. Int. J. Clin. Pharmacol. 22, 63. Saez-Llorens, X., Odio, C.M., Umana, M.A. and Morales, M.V. 1989. Spiramycin vs. placebo for treatment of acute diarrhea caused by Cryptosporidium. Pediatr. Infect. Dis. J. 8, 136–140. Scaglia, M., Atzori, C., Marchetti, G., Orso, M., Maserati, R., Orani, A., Novati, S. and Olliaro, P. 1994. Effectiveness of aminosidine (paromomycin) sulfate in chronic Cryptosporidium diarrhea in AIDS patients: An open, uncontrolled, prospective clinical trial. J. Infect. Dis. 170, 1349–1350. Schmidt, W., Wahnschaffe, U., Schäfer, M., Zippel, T., Arvand, M., Meyerhans, A., Rieken, E.-O. and Ullrich, R. 2001. Rapid increase of mucosal CD4 T cells followed by clearance of intestinal cryptosporidiosis in an AIDS patient receiving highy active antiretroviral therapy. Gastroenterology 120, 984–987. Shepherd, R.C., Smail, P.J. and Sinha, G.P. 1989. Reactive arthritis complicating cryptosporidial infection. Arch. Dis. Child. 64, 743–744. Smith, L.M., Bonafonte, M.-T., Campbell, L.D. and Mead, J.R. 2001. Exogenous interleukin-12 (IL-12) exacerbates Cryptosporidium parvum infection in gamma interferon knockout mice. Exp. Parasitol. 98, 123–133. Smith, N.H., Cron, S., Valdez, L.M., Chappell, C.L. and White, A.C. 1998. Combination drug therapy for cryptosporidiosis in AIDS. J. Infect. Dis. 178, 900–903. Soave, R. 1990. Treatment strategies for cryptosporidiosis. Ann. N. Y. Acad. Sci. 616, 442–451. 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., Dieterich, D., Kotler, D., Gassyuk, E., Tierney, A., Lieber, L. and Legendre, R. 1990. Oral diclazuril for cryptosporidiosis. 6th International Conference on AIDS, Abstract Th.B. 519, San Francisco, CA. Soave, R., Havlir, D., Lancaster, D., Joseph, P., Leedom, J., Clough, W., Geisler, P. and Dunne, M. 1993. Azithromycin therapy of AIDS-related cryptosporidial diarrhea: A multi-center, placebo-controlled, double-blind study. 33rd Interscience Conference on Antimicrobial Agents and Chemotherapy, Abstract 193, New Orleans, LA. Soave, R., Sjoerdsma, A. and Cawein, M.J. 1985. Treatment of cryptosporidiosis in AIDS patients with DFMO. First International Conference on AIDS, Abstract 30, Atlanta. Sprinz, E., Mallman, R., Barcellos, S., Silbert, S., Schestatsky, G. and David, D.B. 1998. AIDS-related cryptosporidial diarrhoea: An open study with roxithromycin. J. Antimicrob. Chemother. 41, 85–91. Sréter, T., Széll, Z. and Varga, I. 1999. Attempted chemoprophylaxis of cryptosporidiosis in chickens, using diclazuril, toltrazuril, or garlic extract. J. Parasitol. 85, 989–991. Sréter, T., Széll, Z. and Varga, I. 2002. Anticryptosporidial prophylactic efficacy of enrofloxacin and paromomycin in chickens. J. Parasitol. 88, 209–211. Sunnotel, O., Lowery, C.J., Moore, J.E., Dooley, J.S.G., Xiao, L., Millar, B.C., Rooney, P.J. and Snelling, W.J. 2006. Under the microscope: Cryptosporidium. Lett. Appl. Microbiol. 43, 7–16. Teixidor, H.S., Godwin, T.S. and Ramirez, E.A. 1991. Cryptosporidiosis of the biliary tract in AIDS. Radiology. 180, 51–56.



287

Tenter, A.M., Barta, J.R., Beveridge, I., Duszynski, D.W., Mehlhorn, H., Morrison, D.A., Thompson, R.C.A. and Conrad, P.A. 2002. The conceptual basis for a new classification of the coccidia. Int. J. Parasitol. 32, 595–616. Thompson, R.C.A., Olson, M.E., Zhu, G., Enomoto, S., Abrahamsen, M.S. and Hijjawi, N.S. 2005. Cryptosporidium and cryptosporidiosis. Adv. Parasitol. 59, 77–158. Trad, O., Jumaa, P., Uduman, S. and Nawaz, A. 2003. Eradication of Cryptosporidium in four children with acute lymphoblastic leukemia. J. Trop. Pediat. 49, 128–130. Tzipori, S.R., Campbell, I. and Angus, K. 1982. The therapeutic effect of 16 antimicrobial agents on Cryptosporidium infection in mice. Aust. J. Exp. Biol. Med. Sci. 60, 187–190. Uip, D.E., Lima, A.L.L., Amato, V.S., Boulos, M., Neto, V.A. and David, D.B. 1998. Roxithromycin treatment for diarrhoea caused by Cryptosporidium spp. in patients with AIDS. J. Antimicrob. Chemother. 41, 93–97. Ungar, B.L.P., 1994. Cryptosporidium and cryptosporidiosis, in Textbook of AIDS Medicine, Broder, S., Merigan, T.C. and Bolognesi, D. (Eds). Williams & Wilkins, Baltimore, MD, 951. 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. Vargas, S.L., Shenep, J.L., Flynn, P.M., Pui, C.H., Santana, V.M. and Hughes, W.T. 1993. Azithromycin for the treatment of severe Cryptosporidium diarrhea in two children with cancer. J. Pediatr. 123, 154–156. Verdon, R., Polianski, J., Gaudebout, C., Marche, C., Garry, L., Carbon, C. and Pocidalo, J.J. 1995. Evaluation of high dose regimen of paromomycin against Cryptosporidium in the dexamethasone-treated rat model. Antimicrob. Agents Chemother. 39, 2155–2157. Verdon, R., Polianski, J., Gaudebout, C., Marche, C., Garry, L. and Pocidalo, J.J. 1994. Evaluation of curative anticryptosporidial activity of paromomycin in a dexamethasone-treated rat model. Antimicrob. Agents Chemother. 38, 1681–1682. Victor, G.H., Conway, B., Hawley-Foss, N.C., Manion, D. and Sahai, J. 1993. Letrazuril therapy for cryptosporidiosis: Clinical response and pharmacokinetics. AIDS 7, 438–440. Villacorta, I., Peeters, J.E., Vanopdenbosch, E., Ares-Mazas, E. and Theys, H. 1991. Efficacy of halofuginone lactate against Cryptosporidium parvum in calves. Antimicrob. Agents Chemother. 35, 283–287. Viu, M., Quílez, J., Sánchez-Acedo, C., Cacho, E.D. and López-Bernad, F. 2000. Field trial on the therapeutic efficacy of paromomycin on natural Cryptosporidium parvum infections in lambs. Vet. Parasitol. 90, 163–170. Wallace, M.R., Nguyen, M.T. and Newton, J.A. 1993. Use of paromomycin for the treatment of cryptosporidiosis in patients with AIDS. Clin. Infect. Dis. 17, 1070–1071. Waters, W.R., Frydman, B., Marton, L.J., Valasinas, A., Reddy, V.K., Harp, J.A., Wannemuehler, M.J. and Yarlett, N. 2000. [1N,12N]Bis(Ethyl)-cis-6,7-Dehydrospermine: A new drug for treatment and prevention of Cryptosporidium parvum infection of mice deficient in T-cell receptor alpha. Antimicrob. Agents Chemother. 44, 2891–2894. Weikel, C., Lazenby, A., Belitsos, P., McDewit, M., Fleming, H. and Barbacci, M. 1991. Intestinal injury associated with spiramycin therapy of Cryptosporidium diarrhea in two children with cancer. J. Protozool. 38, 147S. Weist, P.M., Johnson, J.H. and Flanigan, T.P. 1993. Microtubule inhibitors block Cryptosporidium parvum infection of a human enterocyte cell line. Infect. Immun. 61, 4888–4890. Whiteside, M.E., Barkin, J.S., May, R.G., Weiss, S.D., Fischl, M.A. and Macleod, C.L. 1984. Enteric coccidiosis among patients with the acquired immunodeficiency syndrome. Am. J. Trop. Med. Hyg. 33, 1065–1072. Wittenberg, D.F., Miller, N.M. and Ende, J.V.D. 1989. Spiramycin is not effective in treating Cryptosporidium diarrhea in infants: Results of a double-blind randomized trial. J. Infect. Dis. 159, 131–132. 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. Woolf, G.M., Townsend, M. and Guyatt, G. 1987. Treatment of cryptosporidiosis with spiramycin in AIDS. An “N of 1” trial. J. Clin. Gastroenterol. 9, 632–634. Zhanel, G.G., Dueck, M., Hoban, D.J., Vercaigne, L.M., Embil, J.M., Gin, A.S. and Karlowsky, J.A. 2001. Review of macrolides and ketolides: Focus on respiratory tract infections. Drug. 61, 443–498.



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


290


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


Ostrea edulis

Netherland Spain

Crassostrea gigas Cerastoderma edule


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


Foodborne Transmission

291

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).


292


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).



293

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%.


294


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



295

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


296


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



297

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


298


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.



299

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


300


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.

References Anonymous. 1984. Cryptosporidiosis surveillance. Wkly. Epidemiol. Rec. 59, 72–73. Anonymous. 1985. A case of food poisoning caused by Cryptosporidium—England. Canada Diseases Weekly Report 11, 76–77. Anonymous. 1996. Foodborne outbreak of diarrheal illness associated with Cryptosporidium parvum—Minnesota, 1995. Morbid. Mortal. Wkly. Rep. 45, 783–784. Anonymous. 1997. Outbreaks of Escherichia coli O157:H7 infection and cryptosporidiosis associated with drinking unpasteurized apple cider—Connecticut and New York, October 1996. Morbid. Mortal. Wkly. Rep., 46, 4–8. Anonymous. 1998. Foodborne outbreak of Cryptosporidiosis—Spokane, Washington, 1997. Morbid. Mortal. Wkly. Rep. 47, 565–567. Anonymous. 1998b. Guide to minimize microbial food safety hazards for fresh fruits and vegetables. Guidance for Industry. U.S. Department of Health and Human Services Food and Drug Administration Center for Food Safety and Applied Nutrition (CFSAN) October, 1998. p. 43. Anonymous. 2004. Preliminary FoodNet Data on the Incidence of Infection with Pathogens Transmitted Commonly Through Food—Selected Sites, United States, 2003. Morbid. Mortal. Wkly. Rep. 53, 338–343 Anonymous. 2005. Preliminary FoodNet Data on the Incidence of Infection with Pathogens Transmitted Commonly Through Food—10 Sites, United States, 2004. Morbid. Mortal. Wkly. Rep. 54, 352–356. Anonymous. 2006. Preliminary FoodNet Data on the Incidence of Infection with Pathogens Transmitted Commonly Through Food—10 States, United States, 2005. Morbid. Mortal. Wkly. Rep. 55, 392–395 Armon, R., Gold, D., Brodsky, M., and Oron, G. 2002. Surface and subsurface irrigation with effluents of different qualities and presence of Cryptosporidium oocysts in soil and on crops. Water Sci. Technol. 46, 115–122. Ashokkumar, M., Vu, T., Grieser, F., Weerawardena, A., Anderson, N., Pilkington, N., and Dixon, D.R. 2003. Ultrasonic treatment of Cryptosporidium oocysts. Water Sci. Technol. 47, 173–177. Bier, J.W. 1991. Isolation of parasites on fruits and vegetables. Southeast Asian J. Trop. Med. Pub. Health 22 Suppl., 144–145. Blackburn, B.G., Mazurek, J.M., Hlavsa, M., Park, J., Tillapaw, M., Parrish, M., Salehi, E., Franks, W., Koch, E., Smith, F., Xiao, L., Arrowood, M., Hill, V., da Silva, A., Johnston, S., Jones, J.L. 2006. Cryptosporidiosis associated with ozonated apple cider. Emerg. Infect. Dis. 12, 684–686. Calvo, M., Carazo, M., Arias, M.L., Chaves, C., Monge, R., and Chinchilla, M. 2004. [Prevalence of Cyclospora sp., Cryptosporidium sp, microsporidia and fecal coliform determination in fresh fruit and vegetables consumed in Costa Rica]. Arch. Latinoam. Nutr. 54, 428–432. Casemore, D.P., Jessop, E.G., Douce, D., and Jackson, F.B. 1986. Cryptosporidium plus Campylobacter: An outbreak in a semi-rural population. J. Hyg. (London) 96, 95–105. Chalmers, R.M., Sturdee, A.P., Mellors, P., Nicholson, V., Lawlor, F., Kenny, F., and Timpson, P. 1997. Cryptosporidium parvum in environmental samples in the Sligo area, Republic of Ireland: A preliminary report. Lett. Appl. Microbiol. 25, 380–384.



301

Chappell, C.L., Okhuysen, PC,, Langer-Curry R,, Widmer, G., Akiyoshi, D.E., Tanriverdi, S., and Tzipori, S. 2006. Cryptosporidium hominis: Experimental challenge of healthy adults. Am. J. Trop. Med. Hyg. 75, 851–7. Conrad, P.A., Miller, M.A., Kreuder, C., James, E.R., Mazet, J., Dabritz, H., Jessup, D.A., Gulland, F., Grigg, M.E. 2005. Transmission of Toxoplasma: Clues from the study of sea otters as sentinels of Toxoplasma gondii flow into the marine environment. Int. J. Parasitol. 35, 1155–1168. Cook, N., Paton, C.A., Wilkinson, N., Nichols, R.A., Barker, K., and Smith, H.V. 2006a. Towards standard methods for the detection of Cryptosporidium parvum on lettuce and raspberries. Part 1: Development and optimization of methods. Int. J. Food Microbiol. 109, 215–221. Cook, N., Paton, C.A., Wilkinson, N., Nichols, R.A., Barker, K., and Smith, H.V. 2006b. Towards standard methods for the detection of Cryptosporidium parvum on lettuce and raspberries. Part 2: Validation. Int. J. Food Microbiol. 109, 222–228. Dawson, D.J., Samuel, C.M., Scrannage, V., and Atherton, C.J. 2004. Survival of Cryptosporidium species in environments relevant to foods and beverages. J. Appl. Microbiol. 96, 1222–1229. Deng, M.Q., & Cliver, D.O. 1999. Cryptosporidium parvum studies with dairy products. Int. J. Food Microbiol. 46, 113–121. Deng, M.Q., & Cliver, D.O. 2000. Comparative detection of Cryptosporidium parvum oocysts from apple juice. Int. J. Food Microbiol. 54, 155–162. Elsser, K.A., Moricz, M., and Proctor, E.M. 1986. Cryptosporidium infections: A laboratory survey. Can. Med. Assoc. J. 135, 211–213. Fayer, R., Graczyk, T.K., Lewis, E.J., Trout, J.M., and Farley, C.A. 1998. Survival of infectious Cryptosporidium parvum oocysts in seawater and eastern oysters (Crassostrea virginica) in the Chesapeake Bay. Appl. Environ. Microbiol. 64, 1070–1074. Fayer, R., Lewis, E.J., Trout, J.M., Graczyk, T.K., Jenkins, M.C., Higgins, J., Xiao, L., and Lal, A.A. 1999. Cryptosporidium parvum in oysters from commercial harvesting sites in the Chesapeake Bay. Emerg. Infect. Dis. 5, 706–710. Fayer, R., Trout, J.M., Graczyk, T.K., and Lewis, E.J. 2000. Prevalence of Cryptosporidium, Giardia and Eimeria infections in post-weaned and adult cattle on three Maryland farms. Vet. Parasitol. 93, 103–112. Fayer, R., Trout, J.M., Lewis, E.J., Santin, M., Zhou, L., Lal, A.A., and Xiao, L. 2003. Contamination of Atlantic coast commercial shellfish with Cryptosporidium. Parasitol. Res. 89, 141–145. Fayer, R., Trout, J.M., Lewis, E.J., Xiao, L., Lal, A., Jenkins, M.C., and Graczyk, T.K. 2002. Temporal variability of Cryptosporidium in the Chesapeake Bay. Parasitol. Res. 88, 998–1003. Feng, Y., Ortega, Y., He, G., Das, P., Xu, M., Zhang, X., Fayer, R., Gatei, W., Cama, V., and Xiao, L. 2007. Wide geographic distribution of Cryptosporidium bovis and the deer-like genotype in bovines. Vet. Parasitol. 144, 1–9. Freire-Santos, F., Gomez-Couso, H., Ortega-Inarrea, M.R., Castro-Hermida, J.A., Oteiza-Lopez, A.M., GarciaMartin, O., and Ares-Mazas, M.E. 2002. Survival of Cryptosporidium parvum oocysts recovered from experimentally contaminated oysters (Ostrea edulis) and clams (Tapes decussatus). Parasitol. Res. 88, 130–133. Freire-Santos, F., Oteiza-Lopez, A.M., Vergara-Castiblanco, C.A., Ares-Mazas, E., Varez-Suarez, E., and Garcia-Martin, O. 2000. Detection of Cryptosporidium oocysts in bivalve molluscs destined for human consumption. J. Parasitol. 86, 853–854. Friedman, D.E., Patten, K.A., Rose, J.B., and Barney, M.C. 1997. The potential for Cryptosporidium parvum oocysts survival in beverages associated with contaminated tap water. J. Food Safety 17, 125–132. Gajadhar, A.A., Scandrett, W.B., and Forbes, L.B. 2006. Overview of food- and water-borne zoonotic parasites at the farm level. Rev. Sci. Tech. 25, 595–606. Garcia, L., Henderson, J., Fabri, M., and Oke, M. 2006. Potential sources of microbial contamination in unpasteurized apple cider. J. Food Prot. 69, 137–144. Gelletlie, R., Stuart, J., Soltanpoor, N., Armstrong, R., and Nichols, G. 1997. Cryptosporidiosis associated with school milk. Lancet 350, 1005–1006. Gomez-Bautista, M., Ortega-Mora, L.M., Tabares, E., Lopez-Rodas, V., and Costas, E. 2000. Detection of infectious Cryptosporidium parvum oocysts in mussels (Mytilus galloprovincialis) and cockles (Cerastoderma edule). Appl. Environ. Microbiol. 66, 1866–1870.


302


Gomez-Couso, H., Freire-Santos, F., Amar, C.F., Grant, K.A., Williamson, K., Ares-Mazas, M.E., and McLauchlin, J. 2004. Detection of Cryptosporidium and Giardia in molluscan shellfish by multiplexed nested-PCR. Int. J. Food Microbiol. 91, 279–288. Gomez-Couso, H., Freire-Santos, F., Hernandez-Cordova, G.A., and Ares-Mazas, M.E. 2005. A histological study of the transit of Cryptosporidium parvum oocysts through clams (Tapes decussatus). Int. J. Food Microbiol. 102, 57–62. Gomez-Couso, H., Freire-Santos, F., Martinez-Urtaza, J., Garcia-Martin, O., and Ares-Mazas, M.E. 2003. Contamination of bivalve molluscs by Cryptosporidium oocysts: The need for new quality control standards. Int. J. Food Microbiol. 87, 97–105. Graczyk, T.K., Conn, D.B., Lucy, F., Minchin, D., Tamang, L., Moura, L.N., and DaSilva, A.J. 2004. Human waterborne parasites in zebra mussels (Dreissena polymorpha) from the Shannon River drainage area, Ireland. Parasitol. Res. 93, 385–391. Graczyk, T.K., Farley, C.A., Fayer, R., Lewis, E.J., and Trout, J.M. 1998a. Detection of Cryptosporidium oocysts and Giardia cysts in the tissues of eastern oysters (Crassostrea virginica) carrying principal oyster infectious diseases. J. Parasitol. 84, 1039–1042. Graczyk, T.K., Fayer, R., Cranfield, M.R., and Conn, D.B. 1998b. Recovery of waterborne Cryptosporidium parvum oocysts by freshwater benthic clams (Corbicula fluminea). Appl. Environ. Microbiol. 64, 427–430. Graczyk, T.K., Fayer, R., Lewis, E.J., Trout, J.M., and Farley, C.A. 1999. Cryptosporidium oocysts in Bent mussels (Ischadium recurvum) in the Chesapeake Bay. Parasitol. Res. 85, 518–521. Graczyk, T.K., Grimes, B.H., Knight, R., da Silva, A.J., Pieniazek, N.J., and Veal, D.A. 2003. Detection of Cryptosporidium parvum and Giardia lamblia carried by synanthropic flies by combined fluorescent in situ hybridization and a monoclonal antibody. Am. J. Trop. Med. Hyg. 68, 228–232. Graczyk, T.K., Marcogliese, D.J., de, L.Y., da Silva, A.J., Mhangami-Ruwende, B., and Pieniazek, N.J. 2001. Cryptosporidium parvum oocysts in zebra mussels (Dreissena polymorpha): Evidence from the St. Lawrence River. Parasitol. Res. 87, 231–234. Graczyk, T.K. & Schwab, K.J. 2000. Foodborne infections vectored by molluscan shellfish. Curr. Gastroenterol. Rep. 2, 305–309. Harper, C.M., Cowell, N.A., Adams, B.C., Langley, A.J., and Wohlsen. 2002. Outbreak of Cryptosporidium linked to drinking unpasteurized milk. CID 26, 449–450. Huamanchay, O., Genzlinger, L., Iglesias, M., and Ortega, Y.R. 2004. Ingestion of Cryptosporidium oocysts by Caenorhabditis elegans. J. Parasitol. 90, 1176–1178. Izumi, T., Itoh, Y., Yagita, K., Endo, T., and Ohyama, T. 2004. Brackish water benthic shellfish (Corbicula japonica) as a biological indicator for Cryptosporidium parvum oocysts in river water. Bull. Environ. Contam. Toxicol. 72, 29–37. Jiang, J., Alderisio, K.A., Singh, A., and Xiao, L. 2005. Development of procedures for direct extraction of Cryptosporidium DNA from water concentrates and for relief of PCR inhibitors. Appl. Environ. Microbiol. 71, 1135–1141. Jokipii, L., & Jokipii, A.M. 1986. Timing of symptoms and oocyst excretion in human cryptosporidiosis. New Engl. J. Med. 315, 1643–1647. Kniel, K.E. and Jenkins, M.C. 2005. Detection of Cryptosporidium parvum oocysts on fresh vegetables and herbs using antibodies specific for a Cryptosporidium parvum viral antigen. J. Food Prot. 68, 1093–1096. Korich, D.G., Mead, J.R., Madore, M.S., Sinclair, N.A., and Sterling, C.R. 1990. Effects of ozone, chlorine dioxide, chlorine, and monochloramine on Cryptosporidium parvum oocyst viability. Appl. Environ. Microbiol. 56, 1423–1428. Laberge, I., Ibrahim, A., Barta, J.R., and Griffiths, M.W. 1996. Detection of Cryptosporidium parvum in raw milk by PCR and oligonucleotide probe hybridization. Appl. Environ. Microbiol. 62, 3259–3264. Lowery, C.J., Nugent, P., Moore, J.E., Millar, B.C., Xiru, X., and Dooley, J.S. 2001. PCR-IMS detection and molecular typing of Cryptosporidium parvum recovered from a recreational river source and an associated mussel (Mytilus edulis) bed in Northern Ireland. Epidemiol. Infect. 127, 545–553. MacKenzie, W.R., Hoxie, N.J., Proctor, M.E., Gradus, M.S., Blair, K.A., Peterson, D.E., Kazmierczak, J.J., Addiss, D.G., Fox, K.R., Rose, J.B., and Davis, J.P. 1994. A massive outbreak in Milwaukee of Cryptosporidium infection transmitted through the public water supply. N. Engl. J. Med. 331, 161–167. MacRae, M., Hamilton, C., Strachan, N.J., Wright, S., and Ogden, I.D. 2005. The detection of Cryptosporidium parvum and Escherichia coli O157 in UK bivalve shellfish. J. Microbiol. Meth. 60, 395–401.



303

Mathison, B.A., & Ditrich, O. 1999. The fate of Cryptosporidium parvum oocysts ingested by dung beetles and their possible role in the dissemination of cryptosporidiosis. J. Parasitol. 85, 678–681. Millar, B.C., Finn, M., Xiao, L., Lowery, C.J., Dooley, J.S., and Moore, J.E. 2002. Cryptosporidium in foodstuffs- and emerging aetiological route of human foodborne illness. Trends Food Sci. Tech.13, 168–187. Millard, P.S., Gensheimer, K.F., Addiss, D.G., Sosin, D.M., Beckett, G.A., Houck-Jankoski, A., and Hudson, A. 1994. An outbreak of cryptosporidiosis from fresh-pressed apple cider. J. Am. Med. Assoc. 272, 1592–1596. Monge, R., & Arias, M.L. 1996. [Presence of various pathogenic microorganisms in fresh vegetables in Costa Rica]. Arch. Latinoam. Nutr. 46, 292–294. Monge, R., Chinchilla, M., and Reyes, L. 1996. [Seasonality of parasites and intestinal bacteria in vegetables that are consumed raw in Costa Rica]. Rev. Biol. Trop. 44, 369–375. Olson, M.E., Goh, J., Phillips, M., Guselle, N., and McAllister, T.A. 2007. Giardia cyst and Cryptosporidium oocyst survival in water, soil and cattle feces. J. Environ. Qual. 28, 1991–1996. Ong, C., Moorehead, W., Ross, A., and Isaac-Renton, J. 1996. Studies of Giardia spp. and Cryptosporidium spp. in two adjacent watersheds. Appl. Environ. Microbiol. 62, 2798–2805. Ortega, Y.R., Roxas, C.R., Gilman, R.H., Miller, N.J., Cabrera, L., Taquiri, C., and Sterling, C.R. 1997. Isolation of Cryptosporidium parvum and Cyclospora cayetanensis from vegetables collected in markets of an endemic region in Peru. Am. J. Trop. Med. Hyg. 57, 683–686. Peeters, J.E., Mazas, E.A., Masschelein, W.J., Villacorta Martiez, D.M., I, and Debacker, E. 1989. Effect of disinfection of drinking water with ozone or chlorine dioxide on survival of Cryptosporidium parvum oocysts. Appl. Environ. Microbiol. 55, 1519–1522. Peng, M.M., Xiao, L., Freeman, A.R., Arrowood, M.J., Escalante, A.A., Weltman, A.C., Ong, C.S., MacKenzie, W.R., Lal, A.A., and Beard, C.B. 1997. Genetic polymorphism among Cryptosporidium parvum isolates: Evidence of two distinct human transmission cycles. Emerg. Infect. Dis. 3, 567–573. Pepin, M., Russo, P., and Pardon, P. 1997. Public health hazards from small ruminant meat products in Europe. Rev. Sci. Tech. 16, 415–425. Quiroz, E.S., Bern, C., MacArthur, J.R., Xiao, L., Fletcher, M., Arrowood, M.J., Shay, D.K., Levy, M.E., Glass, R.I., and Lal, A. 2000. An outbreak of cryptosporidiosis linked to a foodhandler. J. Infect. Dis. 181, 695–700. Ripabelli, G., Leone, A., Sammarco, M.L., Fanelli, I., Grasso, G.M., and McLauchlin, J. 2004. Detection of Cryptosporidium parvum oocysts in experimentally contaminated lettuce using filtration, immunomagnetic separation, light microscopy, and PCR. Foodborne. Pathog. Dis. 1, 216–222. Robertson, L.J., Campbell, A.T., and Smith, H.V. 1992. Survival of Cryptosporidium parvum oocysts under various environmental pressures. Appl. Environ. Microbiol. 58, 3494–3500. Robertson, L.J., & Gjerde, B. 2001a. Factors affecting recovery efficiency in isolation of Cryptosporidium oocysts and Giardia cysts from vegetables for standard method development. J. Food Prot. 64, 1799–1805. Robertson, L.J., & Gjerde, B. 2001b. Occurrence of parasites on fruits and vegetables in Norway. J. Food Prot. 64, 1793–1798. Robertson, L.J., & Gjerde, B. 2000. Isolation and enumeration of Giardia cysts, Cryptosporidium oocysts, and Ascaris eggs from fruits and vegetables. J. Food Prot. 63, 775–778. Robertson, L.J., Johannessen, G.S., Gjerde, B.K., and Loncarevic, S. 2002. Microbiological analysis of seed sprouts in Norway. Int. J. Food Microbiol. 75, 119–126. Rose, J.B. 1997. Environmental ecology of Cryptosporidium and public health implications. Ann. Rev. Pub. Health 18, 135–161. Santin, 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. Schets, F.M., van den Berg, H.H., Engels, G.B., Lodder, W.J., and de Roda Husman, A.M. 2007. Cryptosporidium and Giardia in commercial and non-commercial oysters (Crassostrea gigas) and water from the Oosterschelde, the Netherlands. Int. J. Food Microbiol. 113, 189–194. 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. Shuval, H.I. 1991. Effects of wastewater irrigation of pastures on the health of farm animals and humans. Rev. Sci. Tech. 10, 847–866.


304


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. Prev. Vet. Med. 43, 253–267. Smith, H., & Nichols, R.A. 2006. Zoonotic protozoa—food for thought. Parasitol. 48, 101–104. Smith, H.V., Brown, J., Coulson, J.C., Morris, G.P., and Girdwood, R.W. 1993. Occurrence of oocysts of Cryptosporidium sp. in Larus spp. gulls. Epidemiol. Infect. 110, 135–143. Smith, H.V., Campbell, B.M., Paton, C.A., and Nichols, R.A. 2002. Significance of enhanced morphological detection of Cryptosporidium sp. oocysts in water concentrates determined by using 4′,6′-diamidino-2phenylindole and immunofluorescence microscopy. Appl. Environ. Microbiol. 68, 5198–5201. Szostakowska, B., Kruminis-Lozowska, W., Racewicz, M., Knight, R., Tamang, L., Myjak, P., and Graczyk, T.K. 2004. Cryptosporidium parvum and Giardia lamblia recovered from flies on a cattle farm and in a landfill. Appl. Environ. Microbiol. 70, 3742–3744. Tamburrini, A. & 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. Thurston-Enriquez, J.A., Watt, P., Dowd, S.E., Enriquez, R., Pepper, I.L., and Gerba, C.P. 2002. Detection of protozoan parasites and microsporidia in irrigation waters used for crop production. J. Food Prot. 65, 378–382. Traversa, D., Giangaspero, A., Molini, U., Iorio, R., Paoletti, B., Otranto, D., and Giansante, C. 2004. Genotyping of Cryptosporidium isolates from Chamelea gallina clams in Italy. Appl. Environ. Microbiol. 70, 4367–4370. U.S. Environmental Protection Agency. 2001. Method 1623: Cryptosporidium and Giardia in water by filtration/IMS/FA. Publication No. EPA-821-R-99-006. Office of Water, USEPA. 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. Compost Sci. Util. 12, 235–241. Xiao, L., Sulaiman, I., Fayer, R., and Lal, A.A. 1998. Species and strain-specific typing of Cryptosporidium parasites in clinical and environmental samples. Mem. Inst. Oswaldo Cruz 93, 687–691. Yu, J.R., & Park, W.Y. 2003. The effect of gamma-irradiation on the viability of Cryptosporidium parvum. J. Parasitol. 89, 639–642.

52268_C011.fm Page 305 Friday, August 31, 2007 12:21 PM

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

305


306


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


Waterborne: Drinking Water

307

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


308


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



309

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.


310


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



311

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


312


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



313

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


314


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



315

(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


316


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 1.0c 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


320


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



321

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.


322


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


(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.



323

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.


324


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

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



325

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


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.


326


References Aboytes, R., DiGiovanni, G.D., Abrams, F.A., and LeChevallier, M.W. 2004. Detection of infectious Cryptosporidium in filtered drinking water. JAWWA 96, 88–98. Aguilar, Z.P. and Fritsch, I. 2003. Immobilized enzyme-linked DNA-hybridization assay with electrochemical detection for Cryptosporidium parvum hsp70 mRNA. Anal. Chem. 75, 3890–3897. Aldom, J.E. and Chagla A.H. 1995. Recovery of Cryptosporidium oocysts from water by a membrane filtration dissolution method. Lett. Appl. Microbiol. 20, 186–187. Aldom, J.E. and Chagla, A.H. 1997. Membrane filter dissolution, a novel method for the recovery of Cryptosporidium oocysts from water, in Proc. 1997 Int. Symp. Waterborne Cryptosporidium, Fricker, C.R., Clancy, J.L. and Rochelle, P., Eds., AWWA, Denver, CO, 87–90. Anon. 1991. Proposed test method for Giardia cysts and Cryptosporidium oocysts in low-turbidity water by a fluorescent antibody procedure, in Annual Book ASTM Standards. American Society for Testing and Materials. Philadelphia, PA, 925–935. Anon. 2005. Outbreak of cryptosporidiosis in Northwest Wales, 2005. http://www.hpa.org.uk/cdr/archives/ 2006/cdr2806.pdf. Arrowood, M.J., and Sterling C.R. 1989. Comparison of conventional staining methods and monoclonal antibody-based methods for Cryptosporidium oocyst detection, J. Clin. Microbiol. 27, 1490–1495. Atherholt, T.B. and Korn, L. 1999. ICR protocol, alternative treatment of parasite sample data, JAWWA 91, 95–102. Borchardt, M.A. and Spencer, S.K. 2002. Concentration of Cryptosporidium, microsporidia and other waterborne pathogens by continuous separation channel centrifugation. J. Appl. Microbiol. 92, 649–656. Bukhari, Z., McCuin, R.M., Fricker, C.R., and Clancy, J.L. 1998. Immunomagnetic separation (IMS) of Cryptosporidium parvum from source water samples of various turbidities. Appl. Environ. Microbiol. 64, 4495–4499. Bukhari, Z., Hargy, T.M., Bolton , J.R., Dussert, B., and Clancy, J.L. 1999. Medium-pressure UV light for oocyst inactivation. JAWWA 91, 86–94. Bukhari, Z., Clancy, J.L., Marshall, M., Korich, G., Smith, H.V., O’Grady, J., Sykes, N., and Fricker, C. 1998. in Proc. AWWA WQTC. Denver, CO. Callahan, M.R., Rose, J.B., and García-Rubio, L. 2003. Use of multiwavelength transmission spectroscopy for the characterization of Cryptosporidium parvum oocysts, quantitative interpretation. Environ. Sci. Technol. 37, 5254–5261. Campbell, A.T., Robertson, L.J., Snowball, M.R., and Smith, H.V. 1995. Inactivation of oocysts of Cryptosporidium parvum by ultraviolet irradiation. Water Res. 29, 2583–2586. Campbell, A.T., Grøn, B., and Johnson, S.E. 1997. Immunomagnetic separation of Cryptosporidium oocysts from high turbidity water sample concentrates. in Proc. 1997 Int. Symp. Waterborne Cryptosporidium. Fricker, C.R., Clancy, J.L., and Rochelle, P., Eds. AWWA. Denver, CO. 91–96. Carmena, D., Aguinagalde, X., Zigorraga, C., Fernández-Crespo, J.C., and Ocio, J.A. 2007. Presence of Giardia cysts and Cryptosporidium oocysts in drinking water supplies in northern Spain. J. Appl. Microbiol. 102, 619–629. Chauret, C., Springthorpe, V.S., and Sattar, S.A. 1995. Detection and survival of protozoan parasites in water in the Ottawa (Canada) region, in Protozoan Parasites in Water. Betts, W.B., Casemore, D., Fricker, C., Smith, H., and Watkins, J. Eds. The Royal Society of Chemistry. Cambridge. 81–83. Clancy, J.L., Gollnitz, W.D., and Tabib, Z. 1994. Commercial labs, how accurate are they? JAWWA 86, 89–97. Clancy, J.L. 1996. Development of performance evaluation (PE) sample preparation protocols for Giardia cysts and Cryptosporidium oocysts. USEPA Contract No. 68-C3-0365. Clancy, J.L., Hargy, T.M., and Schaub, S. 1997. Improved sampling methods for the recovery of Giardia and Cryptosporidium from source and treated water, in Proc. 1997 Int. Symp. Waterborne Cryptosporidium. Fricker, C.R., Clancy, J.L., and Rochelle, P. Eds. AWWA. Denver, CO, 79–86. Clancy, J.L., Hargy, T.M., Marshall, M.M., and Dyksen, J.E. 1998. Inactivation of Cryptosporidium parvum oocysts in water using ultraviolet light. JAWWA 90, 92–102. Clancy, J.L., Bukhari, Z., McCuin, R.M., Matheson, Z., and Fricker, C.R. 1999. USEPA method 1622. JAWWA 91, 60–67.



327

Clancy, J.L., Bukhari, Z., McCuin, R.M. Hargy, T.M., Fricker, C.R., Matheson, Z., and Sykes, N. 2000a. New approaches for isolation of Cryptosporidium and Giardia. AwwaRF and AWWA. Denver, CO. Clancy, J.L. 2000b. Sydney’s 1998 water quality crisis. JAWWA 92, 55–66. Clancy, J.L., Bukhari, Z., McCuin, R.M., Clancy, T.P., Marshall, M.M., Korich, D.G., Fricker, C.R., Sykes, N., Smith, H.V., O’Grady, J., Rosen, J.S., Sobrinho, J., and Schaefer, III, F.W. 2000c. Cryptosporidium viability and infectivity methods. AwwaRF and AWWA. Denver, CO. Clancy, J.L., Bukhari, Z., Hargy, T.M. Bolton, J.R., Dussert, B., and Marshall, M.M. 2000d. Using UV to inactivate Cryptosporidium. JAWWA 92, 97–104. Clancy, J.L., Hargy, T.M., Marshall, M.M., Korich, D.G., and Nicholson, W.L. 2002. Susceptibility of multiple strains of C. parvum oocysts to UV light. AwwaRF and AWWA, Denver, CO. Clancy, J.L., Connell, K., and McCuin, R.M. 2003. Implementing PBMS improvements to USEPA’s Cryptosporidium and Giardia methods. JAWWA 95, 80–93. Clancy, J.L. and P.R. Hunter. 2004. Monitoring Giardia and Cryptosporidium in water in the UK and US. 2004, in World Class Parasites, C.R. Sterling, and R.D. Adam, Eds. 129–139. Kluwer Academic Press, Dordrecht. Colbourne, J. 1989. Thames Water Utilities experience with Cryptosporidium, in Proc. AWWA Water Qual. Technol. Conf. AWWA. Denver, CO. Craik, S.A., Weldon, D., Finch, G.R., Bolton, J.R., and Belosevic, M. 2001. Inactivation of Cryptosporidium parvum oocysts using medium- and low-pressure ultraviolet radiation. Water Res. 35, 1387–1398. Current, W.L. and Haynes, T.B. 1984. Complete development of Cryptosporidium in cell culture. Science 22, 603–605. DiGiovanni, G.D., Hashemi, F.H., Shaw, N.J., Abrams, F.A. LeChevallier, M.W., and Abbaszadegan, M. 1999. Detection of infectious Cryptosporidium parvum oocysts in surface and filter backwash water samples by immunomagnetic separation and cell culture-PCR. Appl. Environ. Microbiol. 65, 3427–3432. D’Antonio, R.G., Taylor, J.P., Gustafson, T.L., Current, W.L., Rhodes, M.M., Gary, G.W., and Jr. Zajac., R.A. 1985. A waterborne outbreak of cryptosporidiosis in normal hosts. Ann. Intern. Med. 103, 886–888. de Roubin, M.-R., Pharamond, S., Zanelli, F., Poty, F., Houdart, S., Laurent, F., Drocourt, J.-L., and Van Poucke, S. 2002. Application of laser scanning cytometry followed by epifluorescent and differential interference contrast microscopy for the detection and enumeration of Cryptosporidium and Giardia in raw and potable waters. J. Appl. Microbiol. 93, 599–607. Dugan, N., Fox, K., Owens, J., and Miltner, R. 2001. Controlling Cryptosporidium oocysts using conventional treatment. JAWWA 93, 64–76. Dunn, J., Ott, T. and Clark, W. 1995. Pulsed light treatment of food and packaging. Food Technol. 49, 95. Edzwald, J. and Kelly, M. 1998. Control of Cryptosporidium from reservoirs to clarifiers to filters. Water Sci. Technol. 37, 1–8. Emelko, M., Huck, P. Douglas, I.I., and van den Oever, J. 2000. Cryptosporidium and microsphere removal during low turbidity end-of-run and early breakthrough filtration, in Proc. AWWA Water Qual. Technol. Conf., AWWA, Denver, CO. Ferrari, B. C., et al., G. Vesey, C. Weir, K. L. Williams, and D. A. Veal. 1999. Comparison of Cryptosporidiumspecific and Giardia-specific monoclonal antibodies for monitoring water samples. Water Res. 33, 1611–1617. Finch, G.R., Black, E.K., Gyurek, L., and Belosevic, M. 1993. Ozone inactivation of Cryptosporidium parvum in demand-free phosphate buffer determined by in vitro excystation and animal infectivity. Appl. Environ. Microbiol. 59, 4203-4210. Finch, G.R., Gyurek, L.L., Liyanage, L.R.J., and Belosevic, M. 1997. Effects of various disinfection methods on the inactivation of Cryptosporidium. AWWA and AwwaRF. Denver, CO. Finch, G.R. and M. Belosevic, 1999. Inactivation of Cryptosporidium parvum and Giardia muris with medium pressure ultraviolet radiation, in Proc. USEPA Workshop on UV Disinfection of Drinking Water. Arlington, VA. Fricker, C.R., Jonas, A., Crabb, J., Turner, N., and Smith, H.V. 1997. The concentration and separation of Cryptosporidium oocysts and Giardia cysts using vortex flow filtration and immunomagnetic separation, in Proc. 1997 Int. Symp. Waterborne Cryptosporidium. Fricker, C.R., Clancy, J.L., and Rochelle, P. Eds. AWWA. Denver, CO, 1–8.


328


Fricker, C.R., Turner, N.B., Rolchigo, P.M., Margolin, A.D., and Crabb, J.H. 1995. Improved recovery of Cryptosporidium oocysts from drinking water using vortex flow filtration combined with immunomagnetic separation. J. Am. Soc. Trop. Med. Hyg., 53, 285. Gates, D.G. 1998. The Chlorine Dioxide Handbook. Water Disinfection Series. AWWA. Denver, CO, chap. 1. Graczyk, T.K., Cranfield, M.R., and Fayer, R. 1997. Recovery of waterborne oocysts of Cryptosporidium from water samples by the membrane-filter dissolution method. Parasitol. Res. 83, 121–125. Gregory, R. and Zabel, T. 1999. Sedimentation and flotation, in Water Quality and Treatment, a Handbook of Community Water Supplies, 4th ed. Pontius, F.W., Ed. AWWA. New York. McGraw Hill, 367–450. Grimason, A.M., Smith, H.V., Parker, J.F.W., Bukhari, Z., Campbell, A.T., and Robertson, L.J. 1994. Application of DAPI and immunofluorescence for enhanced identification of Cryptosporidium spp. oocysts in water samples. Water Res. 28, 733–736. Guy, R.A., Payment, P., Krull, U.J., and Horgen, P.A. 2003. Real-time PCR for quantification of Giardia and Cryptosporidium in environmental water samples and sewage. Appl. Environ. Microbiol. 69, 5178–5185. Hall, T., Pressdee, J., and Carrington. E., Removal of Cryptosporidium oocysts by water treatment processes, Report FR0457, Foundation for Water Research, Bucks, U.K., 1994. Hancock, C.M., Rose, J.B., and Callahan, M. 1998. Crypto and Giardia in U.S. groundwater. JAWWA 90, 58–61. Harrington, G., Chen, JH., Harris, A., Xagoraraki, I.I., Battigelli, D., and Standridge, J. 2001. Removal of emerging waterborne pathogens, AwwaRF and AWWA, Denver, CO. Hashimoto, A., Kunikane, S., and Hirata, T. 2002. Prevalence of Cryptosporidium oocysts and Giardia cysts in the drinking water supply in Japan. Water Res. 36, 519–526. Hayes, E.B., Matte, T.D., O’Brien, T.R., McKinley, T.W., Logsdon, G.S., Rose, J.B., Ungar, B.L.P., Word, D.M., Pinsky, P.F., Cummings, M.L., Wilson, M.A., Long, E.G., Hurwitz, E.S., and Juranek, D.D. 1989. Large community outbreak of cryptosporidiosis due to contamination of a filtered public water supply. New Engl. J. Med. 320, 1372–1376. Higgins, J.A., Higgins, J.A., Trout, J.M., Fayer, R., Shelton, D., and Jenkins, C.M. 2003. Recovery and detection of Cryptosporidium parvum oocysts from water samples using continuous flow centrifugation. Water Res. 37, 3551–3560. Ho, B.S.W. and Tan, T.Y. 1998. Giardia and Cryptosporidium in sewage and contaminated river waters. Water Res. 32, 2860–2864. Hoffman, R.M., Standridge, J.H., Chauret, C., and Peterson, L. 1999. Evaluation of four commercially antibodies. JAWWA 91, 69–78. Hörman, A., Rimhanen-Finne, R., Maunula, L., von Bonsdorff, C.H., Torvela, N., Heikinheimo, A., and Hanninen, M.L. 2004. Campylobacter spp., Giardia spp., Cryptosporidium spp., noroviruses, and indicator organisms in surface water in southwestern Finland, 2000–2001. Appl. Environ. Microbiol. 70, 87–95. Hsu, B.-M., Huang, C., Hsu, C.L., Hsu, Y.F., and Yeh, J.H. 1999. Occurrence of Giardia and Cryptosporidium in the Kau-Ping River and its watershed in southern Taiwan. Water Res. 33, 2701. Hsu, B-M., Huang, C., Hsu, Y-F., Jiang, G-Y., and Hsu, C.L. 2001. Evaluation of two concentration methods for detecting Giardia and Cryptosporidium in water. Water Res. 35, 419–429. Huck, P.M., Coffey, B., O’Melia, C., and Emelko, M. 2000. Removal of Cryptosporidium by filtration under conditions of process challenge. in Proc. AWWA Water Qual. Technol. Conf., AWWA. Denver, CO. Humphreys, S., McCreadie, S. and Smith, H.V. 1995. The occurrence and viability of Cryptosporidium sp. oocysts in an upland raw water source in central Scotland, in Protozoan Parasites in Water. Betts, W.B., Casemore, D., Fricker, C., Smith, H., and Watkins, J. Eds., The Royal Society of Chemistry. Cambridge, 87–90. Hunter, P.R. 2000. Advice on the response to reports from public and environmental health to the detection of cryptosporidial oocysts in treated drinking water. Commun. Dis. Pub. Health 3, 24–27. Hutton, P., Ashbolt, N., Vesey, G., Walker, J., and Ongerth, J. 1995. Cryptosporidium and Giardia in the aquatic environment of Sydney, Australia, in Protozoan Parasites in Water. Betts, W.B., Casemore, D., Fricker, C., Smith, H., and Watkins, J. Eds., The Royal Society of Chemistry, Cambridge, 71–75. Jakubowski, W. and Erickson, T.H. 1979. Methods for detection of Giardia cysts in water supplies, in Waterborne Transmission of Giardiasis, Jakubowski, W. and Hoff, J.C., Eds., US Environmental Protection Agency, 600/9-79-001, Cincinnati, OH, 193.



329

Johnson, D.W., Pieniazek, N.J., and Rose, J.B. 1993. DNA probe hybridization and PCR detection of Cryptosporidium compared to immunofluorescence assay. Water Sci. Technol. 27, 77–84. Karanis, P., Maier, W.A., Seitz, H.M., and Schoenen, D. 1992. UV Sensitivity of protozoan parasites. J. Water Supp. Res. Technol.-Aqua. 4, 95–100. Karanis, P., Schoenen, D., and. Seitz, H. M. 1998. Distribution and removal of Giardia and Cryptosporidium in water supplies in Germany. Water Sci. Technol. 37, 9–18. Karanis, P. and Kimura, A. 2002. Evaluation of three flocculation methods for the purification of Cryptosporidium parvum oocysts from water samples. Lett. Appl. Microbiol. 34, 444–449. Kaucner, C. and Stinear, T. 1998. Sensitive and rapid detection of viable Giardia cysts and Cryptosporidium parvum oocysts in large-volume water samples with wound fiberglass cartridge filters and reverse transcription–PCR. Appl. Environ. Microbiol. 64, 743–749. Klevens, C.M. 2001. Parameters for UV disinfection of surface water, in Proc. First International Congress on Ultraviolet Technologies. 2001. Washington, D.C. Korich, D.G., Mead, J.R., Madore, N.A., and Sterling, C.R. 1990. Effects of ozone, chlorine dioxide, chlorine, and monochloramine on Cryptosporidium parvum oocyst viability. Appl. Environ. Microbiol. 56, 1423–1428. Kozwich, D., Johansen, K.A., Landau, K., Roehl, C.A., Woronoff, S., and Roehl, P.A. 2000. Development of a novel, rapid integrated Cryptosporidium parvum detection assay. Appl. Environ. Microbiol. 66, 2711–2717. Kuhn, R.C. and Oshima, K.H. 2001. Evaluation and optimization of a reusable hollow fiber ultrafilter as a first step in concentrating Cryptosporidium parvum oocysts from water, Water Res. 35, 2779–2783. Kuhn, R.C. and Oshima, K.H. 2002. Hollow-fiber ultrafiltration of Cryptosporidium parvum oocysts from a wide variety of 10-L surface water samples. Can. J. Microbiol. 48, 542–549. LaFrenz, R. 1997. High intensity pulsed UV for drinking water treatment, in Proc. AWWA WQTC, Denver, CO. LeChevallier, M.W., Trok, T.O., Burns, M.O., and Lee, R.G. 1990. Comparison of zinc sulfate and immunofluorescence techniques for detecting Giardia and Cryptosporidium in water. JAWWA 82, 75–82. LeChevallier, M.W., Norton, W.D., and Lee, R.G. 1991a. Occurrence of Giardia and Cryptosporidium spp. in surface water supplies. Appl. Environ. Microbiol. 57, 2610–2616. LeChevallier, M.W., Norton, W.D., and Lee, R.G. 1991b. Giardia and Cryptosporidium spp. in filtered drinking water supplies. Appl. Environ. Microbiol. 57, 2617–2621. LeChevallier, M.W. and Norton, W.D. 1995. Giardia and Cryptosporidium in raw and finished water. JAWWA 87, 54–68. LeChevallier, M.W., Norton, W.D., Siegel, J.E., and Abbaszadegan, M. 1995. Evaluation of the immunofluorescence procedure for detection of Giardia cysts and Cryptosporidium oocysts in water. Appl. Environ. Microbiol. 61, 690–697. 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. Li, H., Finch, G.R., and Belosevic. M. 1999. Effect of water temperature on the sequential inactivation of Cryptosporidium using ozone followed by free chlorine, in Proc. AWWA WQTC, Denver, CO. Li, H., Finch, G., Smith, D., and Belosevic, M. 2001. Sequential disinfection design criteria for inactivation of Cryptosporidium oocysts in drinking water. AWWA and AwwaRF, Denver, CO. Li, S. Goodrich, J., Owens, J., Willeke, G., Schaefer, F., and Clark, R. 1997. Reliability of surrogates for determining Cryptosporidium removal. JAWWA 89, 90–98. Linden, K.G., Shin, G., and Sobsey, M.D. 2001. Comparative effectiveness of UV wavelengths for the inactivation of Cryptosporidium parvum oocysts in water. Water Sci. Technol. 43, 171–174. Lowery, C.J., Millar B. C., Moore J. E., Xu J., Xiao L., Rooney P. J., and Crothers, L. 2001. Occurrence and molecular genotyping of Cryptosporidium spp. in surface waters in Northern Ireland. J. Appl. Microbiol. 91, 774–779. Mackey, E.D., Wright, H.B., Hargy, T.M., Malley, Jr., J., and Cushing R.S. 2002. Comparing Cryptosporidium and MS2 bioassays—implications for comparing UV reactor validation. JAWWA 94, 62–69. MacKenzie, W.R., Hoxie, N.J., Proctor, M.E., Gradus, M.S., Blair, K.A., Peterson, D.E., Kazmierczak, J.J., Addis, D.G., Fox, K.R., Rose, J.B., and Davis, J.P. 1994. A massive outbreak in Milwaukee of Cryptosporidium infection transmitted through the public water supply. N. Engl. J. Med. 331, 161–167.


330


Madore, M.S., Rose, J.B. Gerba, C.P., Arrowood, M.J., and Sterling, C.R. 1987. Occurrence of Cryptosporidium spp. oocysts in sewage effluents and selected surface waters. J. Parasitol. 73, 702–705. Matheson, Z., Hargy, T.M., McCuin, R.M.. Clancy, J.L., and Fricker, C.R. 1998. An evaluation of the Gelman Envirochek® capsule for the simultaneous concentration of Cryptosporidium and Giardia from water. J. Appl. Microbiol. 85, 755–761. McCuin, R.M., Bukhari, Z., and Clancy, J.L. 2000. Recovery and viability of Cryptosporidium parvum oocysts and Giardia intestinalis cysts using the membrane dissolution procedure. Can. J. Microbiol., 46, 700–707. McCuin, R.M. and Clancy, J.L. 2003. Modifications to USEPA methods 1622 and 1623 for detection of Cryptosporidium oocysts and Giardia cysts in water. Appl. Environ. Microbiol. 69, 267–274. McTigue, N.E., LeChevallier, M., Arora, H., and Clancy, J. 1998. National assessment of particle removal by filtration. AWWA and AwwaRF. Denver, CO. 1998. Medema, G., Juhasz-Hoterman, M., and Luitjen, J. 2000. Removal of microorganisms by bank filtration in a gravelsand soil, in Proc. Int. Riverbank Filtr. Conf., Dusseldorf, Julich W. and Schubert, J., Eds., Internationale Arbeitgemeinschaft der Wasserwerke im Rheineinzugsgebiet, Amsterdam. Meltzer, T.H. 1993. High Purity Water Preparation, Tall Oaks Publishing, Littleton, CO, p. 232. Mofidi, A.A., Baribeau, H., Rochelle, P.A., De Leon, R., Coffey, B.M., and Green, J.F. 2001. Disinfection of Cryptosporidium parvum with polychromatic UV light. JAWWA 6, 95–109. Monis, P.T. and Saint, C.P. 2001. Development of a nested-PCR assay for the detection of Cryptosporidium parvum in finished water. Water Res., 35, 1641–1648. Musial, C. E., Arrowood, M.J., Sterling, C.R., and Gerba, C.P. 1987. Detection of Cryptosporidium in water by using polypropylene cartridge filters. Appl. Environ. Microbiol. 53, 687–692. National Cryptosporidium Survey Group, A survey of Cryptosporidium oocysts in surface and groundwaters in the U.K. 1992. J. Inst. Wat. Environ. Manage. 6, 697–703. Nichols, G., et al. (2001). Cryptosporidiosis, A report on the surveillance and epidemiology of Cryptosporidium infection in England and Wales Drinking Water Directorate Contract Number DWI 70/2/201. Nieminski, E.C., Schaefer III, F.W., and Ongerth, J.E. 1995. Comparison of two methods for detection of Giardia cysts and Cryptosporidium oocysts in water. Appl. Environ. Microbiol. 61, 1714–1719. Nieminski, E. and Bellamy, W. 2000. Application of surrogate measures to improve treatment plant performance, AWWA and AwwaRF, Denver, CO. Oda, T., Sakagami, M., Ito, H., Yano, H., Rai, S.K., Kawabata, M., and Uga, S. 2000. Size selective continuous flow filtration method for detection of Cryptosporidium and Giardia. Water Res. 34, 4477–4481. Ong C., Moorehead, W., Ross, A., and Isaac-Renton, J. 1996. Studies of Giardia spp. and Cryptosporidium spp. in two adjacent watersheds. Appl. Environ. Microbiol. 62, 2798–2805. Ongerth, J.E. and Stibbs, H.H. 1987. Identification of Cryptosporidium oocysts in river water. Appl. Environ. Microbiol. 53, 672–676. Ongerth, J. and Hutton, P. 1997. DE filtration to remove Cryptosporidium. JAWWA 89, 39–46. Ongerth, J. and Hutton, P. 2001. Testing of diatomaceous earth filtration for removal of Cryptosporidium oocysts. JAWWA 93, 54–63. Ono, K., Tsuji H., Rai, S.K., Yamamoto, A., Masuda, K., Endo, T., Hotta, H., Kawamura, T., and Uga, S., 2001. Contamination of river water by Cryptosporidium parvum oocysts in western Japan. Appl. Environ. Microbiol. 67, 3832–3836. Oppenheimer, J., Aieta, M., Trussell, R., Jacangelo, J., and Najm, I. 2000. Evaluation of Cryptosporidium inactivation in natural waters. AWWA and AwwaRF, Denver, CO. Owens J., Miltner, R., Rice, E., Dahling, D. Schaefer, F., and Shukairy, H. 2000. Pilot-scale ozone inactivation of Cryptosporidium and other microorganisms in natural water. Ozone Sci. Eng. 22, 501–551. Owens J., Miltner, R., Slifko, T., and Rose, J. 1999. In vitro excystation and infectivity in mice and cell culture to assess chlorine dioxide inactivation of Cryptosporidium oocysts. in Proc. AWWA WQTC. Denver, CO. Parker, J.F.W., Smith, H.V., and Girdwood, R.W.A. 1993. Survey at Loch Lomond to assess the occurrence and prevalence of Cryptosporidium spp. oocysts and their likely impact on human health. Foundation for Water Research Report, No. AR 0409. Parkhurst, D.F. and Stein, D.A. 1998. Determining average concentration of Cryptosporidium and other pathogens in water. Water Sci. Technol. 32, 3424–3429. Patania, N., Jacangelo, J., Cummings, L., Wilczak, A., Riley, K., and Oppenheimer, J. 1995. Optimization of filtration for cyst removal, AWWA and AwwaRF, Denver, CO.



331

Patania, N., Mazounie, P., Bernazeau, F., Maclean, G., and Alla, P. 1999. Removal of Cryptosporidium by contact filtration, the Sydney experience, in Proc. AWWA Annu. Conf., Denver, CO. Payment, P., Berube, A., Perreault, D., Armon, R., and Trudel, M. 1989. Concentration of Giardia lamblia cysts, Legionella pneumophila, Clostridium perfringens, human enteric viruses, and coliphages from large volumes of drinking water. Can. J. Microbiol. 35, 932–935. Payment, P. and Franco, E. 1993. Clostridium perfringens and somatic coliphages as indicators of the efficiency of drinking water treatment for viruses and protozoan cysts. Appl. Environ. Microbiol. 59, 2418–2424. Pett, B., Smith, F., Stendahl, D., and Welker, R. 1993. Cryptosporidiosis outbreak from an operations point of view, Kitchener-Waterloo, Ontario Spring 1993, in Proc. AWWA WQTC. Denver, CO. 1993. Plummer, J.D., Edzwald, J.K., and Kelly, M.B. 1995. Removing Cryptosporidium by dissolved-air flotation. JAWWA 87, 85–95. Ransome, M.E., Whitmore, T.N., and Carrington, E.G. 1993. Effects of disinfectants on the viability of Cryptosporidium parvum oocysts. Water Supply (Amsterdam) 11, 75–89. Rennecker J.L., Mariñas, B.J., Owens, J.H., and Rice, E.W. 1999. Inactivation of Cryptosporidium parvum oocysts with ozone. Water Res. 33, 2481–2488. Reynolds, D.T., Slade, R.B., Sykes, N.J., Jonas, A., and Fricker, C.R. 1999. Detection of Cryptosporidium oocysts in water, techniques for generating precise recovery data. J. Appl. Microbiol. 87, 804–813. Roach, P.D., Olson, M.E., Whutley, G., and Wallis, P.M. 1993. Waterborne Giardia cysts and Cryptosporidium oocysts in the Yukon, Canada. Appl. Environ. Microbiol. 59, 67–73. Rochelle, P., De Leon, R., Stewart, M.H., and Wolfe, R. 1997a. Comparison of primers and optimization of PCR conditions for detection of Cryptosporidium parvum and Giardia lamblia in water. Appl. Environ. Microbiol. 63, 106–114. Rochelle, P.A., Ferguson, D.M., Handojo, T.J., De Leon, R., Stewart, M.H, and Wolfe, R.L. 1997b. An assay combining cell culture with reverse transcriptase-PCR to detect and determine infectivity of waterborne Cryptosporidium parvum. Appl. Environ. Microbiol. 63, 2029–2037. Rochelle, P.A., De Leon, R., Johnson, A.M. Stewart, M.H., and Wolfe, R. 1998. Evaluation of immunomagnetic separation for recovery of infectious Cryptosporidium parvum oocysts from environmental samples. Appl. Environ. Microbiol. 65, 841–845. Rochelle, P.A., Marshall, M.M., Mead, J.R., Johnson, A. M., Korich, D.G., Rosen, J.S., and De Leon, R. 2002. Comparison of in vitro cell culture and a mouse assay for measuring infectivity of Cryptosporidium parvum. Appl. Environ. Microbiol. 68, 3809–3817. Rochelle, P.A., Fallar, D., Marshall, M.M., Montelone, B.A., Upton, S., and Woods, K. 2004. Irreversible UV inactivation of Cryptosporidium spp. despite the presence of UV repair genes. J. Euk. Microbiol. 51, 553–562. Roefer, P.A., Monscvitz, J.T., and Rexing, D. 1996. The Las Vegas cryptosporidiosis outbreak. JAWWA 88, 95–106. Rose, J.B., Landeen, L.K., Riley, K.R., and Gerba, C.P. 1989. Evaluation of immunofluorescence techniques for detection of Cryptosporidium oocysts and Giardia cysts from environmental samples. Appl. Environ. Microbiol. 55, 3189–3196. Rose, J.B., Gerba, C.P., and Jakubowski, W. 1991. Survey of potable water supplies for Cryptosporidium and Giardia. Environ. Sci. Technol. 25, 1393–1400. Ruffell, K., Rennecker, J., and Marinas, B. 2000. Inactivation of Cryptosporidium parvum oocysts with chlorine dioxide. Water Res. 34, 868–876. Sartory, D.P., Parton, A., Parton, A.C., Roberts, J., and Bergmann, K. 1998. Recovery of Cryptosporidium oocysts from small and large volume water samples using a compressed foam filter system. Lett. Appl. Microbiol. 27, 318–322. Schuler, P. and Ghosh, M., 1990. Diatomaceous earth filtration of cysts and other particulates using chemical additives. JAWWA 82, 67–75. Schuler, P. and Ghosh, M. 1991. Slow sand filtration of cysts and other particulates, in Proc. AWWA Ann. Conf. Denver, CO. Shepherd, K.M. and Wyn-Jones, A.P. 1996. An evaluation of methods for the simultaneous detection of Cryptosporidium oocysts and Giardia cysts from water. Appl. Environ. Microbiol. 62, 1317–1322. Shin, G.-W., Linden, K.G., Arrowood, M.J., and Sobsey, M.D. 2001. Low-pressure UV inactivation and DNA repair potential of Cryptosporidium parvum oocysts. Appl. Environ. Microbiol. 67, 3029–3032.


332


Siddon, C.A., Chapman, P.Q., and Rush, B.A. 1992. Evaluation of an enzyme immunoassay kit for detecting Cryptosporidium in faeces and environmental samples. J. Clin. Pathol. 45, 479–482. Simmons, O.D., Sobsey, M.D., Heaney, C.D. Schaefer III, F.W., and Francy, D.S. 2001. Concentration and detection of Cryptosporidium oocysts in surface water samples by Method 1622 using ultrafiltration and capsule filtration. Appl. Environ. Microbiol, 67, 1123–1127. Skerrett, H.E. and Holland, C.V. 2000. The occurrence of Cryptosporidium in environmental waters in the greater area. Water Res. 34, 3755–3760. Slifko, T. R., Friedman, D., Rose, J.B., and Jakubowski, W. 1997. An in vitro method for detecting infectious Cryptosporidium oocysts with cell culture. Appl. Environ. Microbiol. 63, 3669–3675. 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. Water Sci. Technol. 24, 169–172. Smith, J.J., Gunasekera, T.S., Barardi, C.R.M., Veal, D., and Vesey, G. (2004) Determination of Cryptosporidium parvum oocyst viability by fluorescence in situ hybridization using a ribosomal RNA-directed probe, J. Appl. Microbiol. 96, 409–417. Sterling, C.R. and Arrowood, M.J. 1986. Detection of Cryptosporidium sp. infections using a direct immunofluorescent assay. Pediatr. Infect. Dis. 5, S139. Stetzenbach, L.D., Arrowood, M.J., Marshall, M.M., and Sterling C.R. 1988. Monoclonal antibody based immunofluorescence assay for Giardia and Cryptosporidium detection in water samples. Water Sci. Technol. 20, 193–198. Swales, C. and Wright, S. 2000. Evaluation of a continuous flow centrifuge for recovery of Cryptosporidium oocysts from large volume water samples. Water Res. 34, 1962–1966. Thurman, R., Faulkner B., Veal D., Cramer, G., and Meiklejohn, M. 1998. Water quality in rural Australia. J. Appl. Microbiol. 84, 627–632. Timms, S., Slade, J., and Fricker, C. 1995. Removal of Cryptosporidium by slow sand filtration. Water Sci. Technol. 31, 81–84. Tyzzer, E.E. 1912. Cryptosporidium parvum (sp. nov.), a coccidian found in the small l intestine of the common mouse. Arch. Protistenkd. 26, 394–412. USEPA. 1989. National primary drinking water regulations, filtration disinfection, turbidity, Giardia lamblia, Viruses, Legionella, and heterotrophic bacteria. Final Rule. Federal Register 54, 124, 27486. USEPA. 1990. Guidance Manual for compliance with the filtration and disinfection requirements for public water systems using surface water sources. Washington, DC. USEPA. 1996. Information Collection Rule, Microbial Laboratory Manual. Office of Research and Development. EPA/600/R-95/178. Washington, DC. USEPA. 1997. National primary drinking water regulations, monitoring requirements for public drinking water supplies. Final Rule. Federal Register 61, 94, 24354. USEPA. 1998. USEPA Method 1622, Cryptosporidium in water by filtration/IMS/IFA. United States Environmental Protection Agency. Office of Water, Washington, DC EPA 821-R-98-010. USEPA. 2001. USEPA Method 1622, Cryptosporidium in water by filtration/IMS/FA. Office of Water. Washington, DC. EPA 821-R-01-026, 2001. USEPA. 2006a. National primary drinking water regulations, long term 2 enhanced surface water treatment rule. Final Rule. Federal Register 40 CFR Parts 9, 141, and 142. USEPA. 2006b. Ultraviolet disinfection guidance manual. Office of Water EPA. 815-R-06-007. Washington, DC. Veal, D., Vesey, G., Fricker, C.R., Ongerth, J., Le Moenic, S., Champion, A., Rossington, G., and Faulkner, B. 1997. Routine cytometric detection of Cryptosporidium and Giardia, recovery rates and quality control, in Proc. 1997 Int. Symp. Waterborne Cryptosporidium. Fricker, C.R., Clancy, J.L., and Rochelle, P., Eds. AWWA, Denver, CO, 9–20. Vesey, G., Slade, J.S., Byrne, M., Shepherd, K., and Fricker, C.R. 1993a. A new method for the concentration of Cryptosporidium oocysts from water. J. Appl. Bacteriol. 75, 82–86. Vesey, G., Slade, J.S., Byrne, M., Shepherd, K., Dennis, P.J., and Fricker, C.R. 1993b. Routine monitoring of Cryptosporidium oocysts in water using flow cytometry. J. Appl. Bacteriol. 75, 87–90. Vesey, G., Hutton, P., Champion, A., Ashbolt, N., Williams, K. L., Warton, A., and Veal, D. A. 1994. Application of flow cytometric methods for the routine detection of Cryptosporidium and Giardia in water. Cytometry 16, 1–6.



333

Wallis, P. M., Erlandsen, S.L., Isaac-Renton, J.L., Olson, M.E., Robertson, W.J., and van Keulen, H. 1996. Prevalence of Giardia cysts and Cryptosporidium oocysts and characterization of Giardia spp. isolated from drinking water in Canada. Appl. Environ. Microbiol. 62, 2789–2797. Wang, J., Song, R., and Hubbs, S. 2000. Particle removal through riverbank filtration process, in Proc. Int. Riverbank Filtr. Conf., Dusseldorf, Julich W. and Schubert, J., Eds., Internationale Arbeitgemeinschaft der Wasserwerke im Rheineinzugsgebiet, Amsterdam. Water Supply. 1999. (Water Quality) (Amendment) Regulations, SI 1524. Stationery Office, London. Water Supply. 2000 (Water Quality) Regulations, SI 3184. Stationery Office, London. Weintraub, J.M. 2006. Improving Cryptosporidium testing methods, a public health perspective. J. Water Health 04, 23–26. Whitmore, T.N. and Carrington, E.G. 1993. Comparison of methods for recovery of Cryptosporidium from water. Water Science Technol. 27, 69–76. Young, P.L. and Komisar, S.J. 1999. The variability introduced by partial sample analysis to numbers of Cryptosporidium oocysts and Giardia cysts reported under the Information Collection Rule. Water Res. 33, 2660–2668. Zuckerman, U., Gold, D., G. Shelef, G., and Armon, R. 1997. The presence of Giardia and Cryptosporidium in surface waters and effluents in Israel. Water Sci. Technol. 35, 381–384. Zuckerman, U., Armon, R., Tzipori, S., and Gold, D. 1999. Evaluation of a portable differential continuous flow centrifuge for concentration of Cryptosporidium oocysts and Giardia cysts from water. J. Appl. Microbiol. 86, 955–961. Zuckerman, U. and Tzipori, S. 2006. Portable continuous flow centrifugation and method 1623 for monitoring of waterborne protozoa from large volumes of various water matrices. J. Appl. Microbiol. 100, 1220–1227.


52268_C012.fm Page 335 Monday, October 15, 2007 6:53 AM

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


336

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;


Waterborne: Recreational Water

337

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,


338


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).



339

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.


340

Cryptosporidium and Cryptosporidiosis, Second Edition 2500

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



341

(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


342


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


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



343

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


1998

June

Wisconsin

12

Pool

Treated


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


344


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).



345

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


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


346


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


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


PHLS, 2000a, 2000c PHLS, 2000a, 2000c PHLS, 2000a, 2000c PHLS, 2000a, 2000c PHLS, 2000a, 2000c PHLS, 1999a, 1999b PHLS, 1999a, 1999b



347

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


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


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.


348


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%


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.



349

Type of Exposure (n=20,441)

Etiologic Agent (n=20,441)

Treated water 73.7%


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%


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.


350

Cryptosporidium and Cryptosporidiosis, Second Edition 35

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



351

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 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



355

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


356


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.



V.

357

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.


358 B.

Cryptosporidium and Cryptosporidiosis, Second Edition Disinfected Water

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



359

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


360


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



361

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.

References American Academy of Pediatrics. 2006. Prevention of illnesses associated with recreational water use. In Pickering, L.K., Baker, C.J., Long, S.S. and McMillan, J.A., eds. 2006. Red Book: 2006 report of the committee on infectious diseases. 27th ed. Elk Grove Village, IL: American Academy of Pediatrics, 199–200. Baker, M., Russell, N., Roseveare, C., O’Hallahan, J., Palmer, S. and Bichan, A. 1998. Outbreak of cryptosporidiosis linked to Hutt Valley swimming pool. N. Z. Pub. Hlth. Rep. 5, 41–45. Barwick, R.S., Levy, D.A., Craun, G.F., Beach, M.J. and Calderon, R.L. 2000. Surveillance for waterborne disease outbreaks—United States, 1997–1998. MMWR Surveill Summ. 49(No. SS-4), 1–35. Beach, M.J., McNeil, M., Arrowood, M., Kreckman, L., Craig, A., Donaldson, K., Sinor, M., Zambie, J., Kauffman, B., Berry, J., Campbell, T., Galvin, V., Stetler, H., Juranek, D. and Addiss, D. 1996. Cryptosporidium in a waterpark: largest U.S. recreational waterborne outbreak [abstract]. In Proceedings of the 45th Annual Epidemic Intelligence Service (EIS) Conference. Atlanta: Centers for Disease Control and Prevention. Beers, M., Cunliffe, D., Kong, P., Laycock, D. and Zonta, D. 1998. Canberra swimming pool associated cryptosporidiosis outbreak. Health Stream 9, 6–8. Bell, A., Guasparini, R., Meeds, D., Mathias, R.G. and Farley, J.D. 1993. A swimming pool-associated outbreak of cryptosporidiosis in British Columbia. Can. J. Pub. Health. 84, 334–337. Betancourt, W.Q. and Rose, J.B. 2004. Drinking water treatment processes for removal of Cryptosporidium and Giardia. Vet. Parasitol. 126, 219–234. Black, M. and McAnulty, J. 2006. The investigation of an outbreak of cryptosporidiosis in New South Wales in 2005. NSW Pub. Health Bull.17, 76–79. Bonadonna, L., Briancesco, R., Magini, V., Orsini, M. and Romano-Spica, V. 2004. A preliminary investigation on the occurrence of protozoa in swimming pools in Italy. Ann. Ig. 16, 709–719. Bukhari, Z., Clancy, J.L., Hary, T.M., Bolton, J., Dussert, B. and Marshall, M.M., 1999. Medium-pressure UV for oocyst inactivation. J. AWWA 91, 86–94. Cabelli, V.J., Dufour, A.P., Levin, M.A., McCabe, L.J. and Haberman, P.W. 1979. Relationship of microbial indicators to health effects at marine bathing beaches. Am. J. Pub. Health 69, 690–696. Carpenter, C., Fayer, R., Trout, J. and Beach, M.J. 1999. Chlorine disinfection of recreational water for Cryptosporidium parvum. Emerg. Infect. Dis. 5, 579–584. Cartwright, R.Y. and Colbourne, J.S. 2002. Cryptosporidiosis and hotel swimming pools—a multifaceted challenge. Water Sci. Technol. 2(3), 47–54. Casemore, D.P., Wright, S.E. and Coop, R.L. 1997. Cryptosporidiosis—Human and animal epidemiology. In Cryptosporidium and Cryptosporidiosis, ed. R Fayer, 65–92. New York, CRC Press. Casemore, D. 2006. Towards a US national estimate of the risk of endemic waterborne disease—seroepidemiologic studies. J. Water Health. 4(Suppl 2), 121–163.


362


Causer, L.M., Handzel, T., Welch, P., Carr, M., Culp, D., Lucht, R., Mudahar, K., Robinson, D., Neavear, E., Fenton, S., Rose, C., Craig, L., Arrowood, M., Wahlquist, S., Xiao, L., Lee, Y.M., Mirel, L., Levy, D., Beach, M.J., Poquette, G. and Dworkin, M.S. 2006. An outbreak of Cryptosporidium hominis infection at an Illinois recreational waterpark. Epidemiol. Infect. 134, 147–156. CDC. 1990. Swimming-associated cryptosporidiosis—Los Angeles County. MMWR Morb. Mortal. Wkly. Rep. 39, 343–345. CDC. 1994. Cryptosporidium infections associated with swimming pools—Dane County, Wisconsin, 1993. MMWR Morb. Mortal. Wkly. Rep. 43, 561–563. CDC. 1998. Outbreak of cryptosporidiosis associated with a water sprinkler fountain—Minnesota, 1997. MMWR Morb. Mortal. Wkly. Rep. 47, 856–860. CDC. 2000. Outbreak of gastroenteritis associated with an interactive water fountain at a beachside park—Florida, 1999. MMWR Morb. Mortal. Wkly. Rep. 49, 565–568. CDC. 2001a. Protracted outbreaks of cryptosporidiosis associated with swimming pool use—Ohio and Nebraska, 2000. MMWR Morb. Mortal. Wkly. Rep. 50, 406–410. CDC. 2001b. Prevalence of parasites in fecal material from chlorinated swimming pools—United States. MMWR Morb. Mortal. Wkly. Rep. 50, 410–412. CDC. 2001c. Responding to fecal accidents in disinfected swimming venues. MMWR Morb. Mortal. Wkly. Rep. 50, 416–417. CDC. 2001d. Shigellosis outbreak associated with an unchlorinated fill-and-drain wading pool—Iowa, 2001. MMWR Morb. Mortal. Wkly. Rep. 50, 797–800. CDC. 2003. Surveillance data from swimming pool inspections—selected states and counties, United States, May–September 2002. MMWR Morb. Mortal. Wkly. Rep. 52, 513–516. CDC. 2004. Aseptic meningitis outbreak associated with echovirus 9 among recreational vehicle campers—Connecticut, 2003. MMWR Morb. Mortal. Wkly. Rep. 53, 710–713. Chalker R.B. and Blaser, M.J. 1988. A review of human salmonellosis, III. Magnitude of Salmonella infection in the United States. Rev. Infect. Dis. 10, 111–124. Chappell, C.L., Okhuysen, P.C., Sterling, C.R. and DuPont, H.L. 1996. Cryptosporidium parvum: intensity of infection and oocyst excretion patterns in healthy volunteers. J. Infect. Dis. 173, 232–236. Cheung, W.H., Chang, K.C., Hung, R.P. and Kleevens, J.W. 1990. Health effects of beach water pollution in Hong Kong. Epidemiol. Infect. 105, 139–162. Clancy, J.L., Bukhari, Z., Hargy, T.M., Bolton, J.R., Dussert, B.W. and Marshall, M.M. 2000. Using U.V. to inactivate Cryptosporidium. J. AWWA 92, 97–104. Clancy, J.L., Marshall, M.M., Hargy, T.H. and Korich, D.G. 2004. Susceptibility of five strains of Cryptosporidium parvum oocysts to UV light. J. AWWA 96, 84–93. Cordell, R.L. and Addiss, D.G. 1994. Cryptosporidiosis in child care settings: a review of the literature and recommendations for prevention and control. Pediatr. Infect. Dis. J. 13, 310–317. Corona-Vasquez, B., Rennecker, J.L., Driedger, A.M. and Marinas, B.J. 2002a. Sequential inactivation of Cryptosporidium parvum oocysts with chlorine dioxide followed by free chlorine or monochloramine. Water Res. 36, 178–188. Corona-Vasquez, B., Samuelson, A., Rennecker, J.L. and Marinas, B.J., 2002b. Inactivation of Cryptosporidium parvum oocysts with ozone and free chlorine. Water Res. 36, 4053–4063. Coupe, S., Delabre, K., Pouillot, R., Houdart, S., Santillana-Hayat, M. and Derouin, F. 2006. Detection of Cryptosporidium, Giardia and Enterocytozoon bieneusi in surface water, including recreational areas: a one-year prospective study. FEMS Immunol. Med. Microbiol. 47, 351–359. Craik, S.A., Weldon, D., Finch, G.R., Bolton, J.R. and Belosevic, M. 2001. Inactivation of Cryptosporidium parvum oocysts using medium- and low-pressure ultraviolet radiation. Water Res. 35, 1387–1398. Craun, G.F., Calderon, R.L. and Craun, M.F. 2005. Outbreaks associated with recreational water in the United States. Int. J. Environ. Health Res. 15, 243–262. Current, W.L. 1985. Cryptosporidiosis. J. Am. Vet. Med. Assoc. 187, 1334–1338. Dadswell, J.V. 1996. Managing swimming, spa, and other pools to prevent infection. Commun. Dis. Rep. CDR Rev. 6(2), R37–40. D’Angelo, L.J., Hierholzer, J.C., Keenlyside, R.A., Anderson, L.J. and Martone, W.J. 1979. Pharyngoconjunctival fever caused by adenovirus type 4: report of a swimming pool-related outbreak with recovery of virus from pool water. J. Infect. Dis. 140, 42–47.



363

Dietz, V.J. and Roberts, J.M. 2000. National Surveillance for Infection with Cryptosporidium parvum, 1995–1998: what Have We Learned? Pub. Health Rep. 115, 358–363. Dufour, A. 1984. Health effects criteria for fresh recreational waters. EPA-600-1-84-004. Cincinnati, OH, U.S. Environmental Protection Agency. Dufour, A.P., Evans, O., Behymer, T.D. and Cantu, R. 2006. Water ingestion during swimming activities in a pool: a pilot study. J. Water Health. 4, 425–430. DuPont, H.L., Chappell, C.L., Sterling, C.R., Okhuysen, P.C., Rose, J.B. and Jakubowski, W. 1995. The infectivity of Cryptosporidium parvum in healthy volunteers. N. Engl. J. Med. 332, 855–859. Dziuban, E.J., Liang, J.L., Craun, G.F., Hill, V., Yu, P.A., Painter, J., Moore, M.R., Calderon, R.L., Roy, S.L. and Beach, M.J. 2006. Surveillance for waterborne disease and outbreaks associated with recreational water—United States, 2003–2004. MMWR Surveill. Summ. 55(No SS-12), 1–30. Egorov, A., Frost, F., Muller, T., Naumova, E., Tereschenko, A. and Ford, T. 2004. Serological evidence of Cryptosporidium infections in a Russian city and evaluation of risk factors for infections. Ann. Epidemiol. 14, 129–136. EPA. 1986. Bacterial ambient water quality criteria for marine and fresh recreational waters. Cincinnati, OH, National Service Center for Environmental Publications; EPA publication no. 440584002. EPA. 2006. EPA’s BEACH report, 2005 swimming season. EPA 823-F-06-010, June 2006. Accessed August 23, 2006 at http://www.epa.gov/waterscience/beaches/seasons/2005/. Fayer, R. and Nerad, T. 1996. Effects of low temperatures on viability of Cryptosporidium parvum oocysts. Appl. Environ. Microbiol. 62, 1431–1433. Fayer, R., Graczyk, T.K., Lewis, E.J., Trout, J.M. and Farley, C.A. 1998a. Survival of infectious Cryptosporidium parvum oocysts in seawater and eastern oysters (Crassostrea virginica) in the Chesapeake Bay. Appl. Environ. Microbiol. 64, 1070–1074. Fayer, R., Trout, J.M. and Jenkins, M.C. 1998b. Infectivity of Cryptosporidium parvum oocysts stored in water at environmental temperatures. J. Parasitol. 84, 1165–1169. Ferley, J.P., Zmirou, D., Balducci, F., Baleux, B., Fera, P., Larbaigt, G., Jacq, E., Moissonnier, B., Blineau, A. and Boudot, J. 1989. Epidemiological significance of microbiological pollution criteria for river recreational waters. Int. J. Epidemiol. 18, 198–205. Fields, B.S., Haupt, T., Davis, J.P., Arduino, M.J., Miller, P.H. and Butler, J.C. 2001. Pontiac fever due to Legionella micdadei from a whirlpool spa: possible role of bacterial endotoxin. J. Infect. Dis. 184, 1289–1292. Fleisher, J.M., Kay, D., Wyer, M.D. and Godfree, A.F. 1998. Estimates of the severity of illnesses associated with bathing in marine recreational waters contaminated with domestic sewage. Int. J. Epidemiol. 27, 722–726. Fournier, S., Dubrou, S., Liguory, O., Gaussin, F., Santillana-Hayat, M., Sarfati, C., Molina, J.M. and Derouin, F. 2002. Detection of Microsporidia, Cryptosporidia and Giardia in swimming pools: a one-year prospective study. FEMS Immunol. Med. Microbiol. 33, 209–213. Frost, F.J., Muller, T., Calderon, R.L. and Craun, G.F. 2000. A serological survey of college students for antibody to Cryptosporidium before and after the introduction of a new water filtration plant. Epidemiol. Infect. 125, 87–92. Frost, F.J., Muller, T.B., Calderon, R.L. and Craun, G.F. 2004. Analysis of serological responses to Cryptosporidium antigen among NHANES III participants. Ann. Epidemiol. 14, 473–478. Furtado, C., Adak, G.K., Stuart, J.M., Wall, P.G., Evans, H.S. and Casemore, D.P. 1998. Outbreaks of waterborne infectious intestinal disease in England and Wales, 1992–5. Epidemiol. Infect. 121, 109–119. Galmes, A., Nicolau, A., Arbona, A., Gomis, E., Guma, M., Smith-Palmer, A., Hernandez-Pezzi, G. and Soler, P. 2003. Cryptosporidiosis outbreak in British tourists who stayed at a hotel in Majorca, Spain. Eurosurveillance Weekly 7(33), 14/08/2003. http://www.eurosurveillance.org/ew/2003/030814.asp). Gerba, C.P. 2000. Assessment of enteric pathogen shedding by bathers during recreational activity and its impact on water quality. Quant. Microbiol. 2, 55–68. Goodgame, R.W., Genta, R.M., White, A.C. and Chappell, C.L. 1993. Intensity of infection in AIDS-associated cryptosporidiosis. J. Infect. Dis. 167, 704–709. Gregory, R. 2002. Bench-marking pool water treatment for coping with Cryptosporidium. J. Environ. Health Res. 1, 11–18.


364


Greinert, J., Furtado, D., Smith, J., Monte Barardi, C. and Simoes, C. 2004. Detection of Cryptosporidium oocysts and Giardia cysts in swimming pool filter backwash water concentrates by flocculation and immunomagnetic separation. Int. J. Environ. Health Res. 14, 395–404. Gustafson, T.L., Band, J.D., Hutcheson, R.H. Jr. and Schaffner, W. 1983. Pseudomonas folliculitis: an outbreak and review. Rev. Infect. Dis. 5, 1–8. Haile, R.W., Witte, J.S., Gold, M., Cressey, R., McGee, C., Millikan, R.C., Glasser, A., Harawa, N., Ervin, C., Harmon, P., Harper, J., Dermand, J., Alamillo, J., Barrett, K., Nides, M. and Wang, G. 1999. The health effects of swimming in ocean water contaminated by storm drain runoff. Epidemiology 10, 355–363. Havelaar, A.H., Bosman, M. and Borst, J. 1983. Otitis externa by Pseudomonas aeruginosa associated with whirlpools. J. Hyg. (Lond). 90, 489–98. Hellard, M.E., Sinclair, M.I., Fairley, C.K., Andrews, R.M., Bailey, M., Black, J., Dharmage, S.C. and Kirk, M.D. 2000. An outbreak of cryptosporidiosis in an urban swimming pool: why are such outbreaks difficult to detect? Aust. N. Z. J. Pub. Health. 24, 272–275. Henrickson, S.E., Wong, T., Allen, P., Ford, T. and Epstein, P.R. 2001. Marine swimming-related illness: implications for monitoring and environmental policy. Environ. Health Perspect. 109, 645–650. Herwaldt, B.L., Craun, G.F., Stokes, S.L. and Juranek, D.D. 1991. Waterborne-disease outbreaks, 1989–1990. MMWR Surveill Summ. 40(No. SS-3), 1–21. Hlavsa, M., Watson, J.C. and Beach, M.J. 2005. Cryptosporidiosis surveillance, United States, 1999–2002. MMWR Surveill Summ. 54 (1), 1–8. Hunt, D.A., Sebugwawo, S., Edmondson, S.G. and Casemore, D.P. 1994. Cryptosporidiosis associated with a swimming pool complex. Commun. Dis. Rep. CDR Rev. 4(2), R20–22. Hunter, P.R., Hughes, S., Woodhouse, S., Syed, Q., Verlander, N.Q., Chalmers, R.M., Morgan, K., Nichols, G., Beeching, N. and Osborn, K. 2004. Sporadic cryptosporidiosis case-control study with genotyping. Emerg. Infect. Dis. 10, 1241–1249. Ichinohe, S., Fukushima, T., Kishida, K., Sanbe, K., Saika, S. and Ogura, M. 2005. Secondary transmission of cryptosporidiosis associated with swimming pool use. Jpn. J. Infect. Dis. 58, 400–401. Insulander, M., Lebbad, M., Stenstrom, T.A. and Svenungsson, B. 2005. An outbreak of cryptosporidiosis associated with exposure to swimming pool water. Scand. J. Infect. Dis. 37, 354–360. Jernigan, D.B., Hofmann, J., Cetron, M.S., Genese, C.A., Nuorti, J.P., Fields, B.S., Benson, R.F., Carter, R.J., Edelstein, P.H., Guerrero, I.C., Paul, S.M., Lipman, H.B. and Breiman, R. 1996. Outbreak of Legionnaires’ disease among cruise ship passengers exposed to a contaminated whirlpool spa. Lancet. 347, 494–499. Joce, R.E., Bruce, J., Kiely, D., Noah, N.D., Dempster, W.B., Stalker, R., Gumsley, P., Chapman, P.A., Norman, P., Watkins, J., Smith, H.V., Price, T.J. and Watts, D. 1991. An outbreak of cryptosporidiosis associated with a swimming pool. Epidemiol. Infect. 107, 497–508. Jokipii, L., and Jokipii, A.M.M. 1986. Timing of symptoms and oocyst excretion in human cryptosporidiosis. N. Engl. J. Med. 315, 1643–1647. Jones, M., Boccia, D., Kealy, M., Salkin, B., Ferrero, A., Nichols, G. and Stuart, J.M. 2006. Cryptosporidium outbreak linked to interactive water feature, U.K.: importance of guidelines. Euro. Surveill. 11, 126–128. Jones, T.F., McMillian, M.B., Scallan, E., Frenzen , Cronquist, A.B., Thomas, S. and Angulo, F.J. 2007. A population-based estimate of the substantial burden of diarrhoeal disease in the United States; FoodNet, 1996–2003. Epidemiol. Infect. 135, 293–301. Kaplan, J.E., Masur, H. and Holmes, K.K. 2002. Guidelines for preventing opportunistic infections among HIV-infected persons—2002. Recommendations of the U.S. Public Health Service and the Infectious Diseases Society of America. MMWR Recomm Rep. 51(RR-8), 1–52. Karanis, P., Sotiriadou, I., Kartashev, V., Kourenti, C., Tsvetkova, N. and Stojanova, K. 2006. Occurrence of Giardia and Cryptosporidium in water supplies of Russia and Bulgaria. Environ. Res. 102, 260–271. Kebabjian, R.S. 1995. Disinfection of public pools and management of fecal accidents. J. Environ. Health 58, 8–12. Kebabjian, R.S. 2003. Interactive water fountains: the potential for disaster. J. Environ. Health 66, 29–30. Kirkpatrick, A., Nichols, G. and Cartwright, R. 2006. Reducing the risks of transmitting gastrointestinal infections in home paddling pools. Euro. Surveill. 31,11(8), E060831.2. Korich, D.G., Mead, J.R., Madore, M.S., Sinclair, N.A. and Sterling, C.R. 1990. Effects of ozone, chlorine dioxide, chlorine, and monochloramine on Cryptosporidium parvum oocyst viability. Appl. Environ. Microbiol. 56, 1423–1428.



365

Kramer, M.H., Herwaldt, B.L., Craun, G.F., Calderon, R.L. and Juranek, D.D. 1996. Surveillance for waterborne-disease outbreaks—United States, 1993–1994. MMWR Surveill. Summ. 45(No. SS-1), 1–33. Kramer, M.H., Sorhage, F.E., Goldstein, S.T., Dalley, E., Wahlquist, S.P. and Herwaldt, B.L. 1998. First reported outbreak in the United States of cryptosporidiosis associated with a recreational lake. Clin. Infect. Dis. 26, 27–33. Learmonth, J.J., Ionas, G., Pita, A.B. and Cowie, R.S. 2003. Identification and genetic characterization of Giardia and Cryptosporidium strains in humans and dairy cattle in the Waikato Region of New Zealand. Water Sci. Technol. 47(3), 21–26. LeChevallier, M.W. and Norton, W.D. 1995. Giardia and Cryptosporidium in raw and finished water. J. AWWA 87, 54–68. LeChevallier, M.W., Norton, W.D. and Lee, R.G. 1991. Occurrence of Giardia and Cryptosporidium spp. in surface water supplies. Appl. Environ. Microbiol. 57, 2610–2616. Lee, S.H., Levy, D.A., Craun, G.F., Beach, M.J. and Calderon, R.L. 2002. Surveillance for waterborne disease outbreaks—United States, 1999–2000. MMWR Surveill. Summ. 51(No. SS-8), 1–47. Lemmon, J.M., McAnulty, J.M. and Bawden-Smith, J. 1996. Outbreak of cryptosporidiosis linked to an indoor swimming pool. Med. J. Aust. 165, 613–616. Levy, D.A., Bens, M.S., Craun, G.F., Calderon, R.L. and Herwaldt, B.L. 1998. Surveillance for waterbornedisease outbreaks—United States, 1995–1996. MMWR Surveill. Summ. 47(No. SS-5), 1–34. Lim, L.S., Varkey, P., Giesen, P. and Edmonson, L. 2004. Cryptosporidiosis outbreak in a recreational swimming pool in Minnesota. J. Environ. Health. 67, 16–20. Louie, K., Gustafson, L., Fyfe, M., Gill, I., MacDougall, L., Tom, L., Wong, Q. and Isaac-Renton, J. 2004. An outbreak of Cryptosporidium parvum in a Surrey pool with detection in pool water sampling. Commun. Dis. Rep. 30(7), 61–66. Macey, J., Lior, L., Johnston, A., Elliott, L., Krahn, D., Nowicki, D. and Wylie, J. 2002. Outbreak of diarrheal illness in attendees at a Ukrainian dance festival, Dauphin, Manitoba—May 2001. Can. Commun. Dis. Rep. 28(17), 141–145. MacKenzie, W.R., Hoxie, N.J., Proctor, M.E., Gradus, M.S., Blair, K.A., Peterson, D.E., Kazmierczak, J.J., Addiss, D.G., Fox, K.R., Rose, J.B. and Davis, J.P. 1994. A massive outbreak in Milwaukee of Cryptosporidium infection transmitted through the public water supply. N. Engl. J. Med. 331, 161–167. MacKenzie, W.R., Kazmierczak, J.J. and Davis, J.P. 1995. An outbreak of cryptosporidiosis associated with a resort swimming pool. Epidemiol. Infect. 115, 545–553. Macro International, Inc. 2000. Cryptosporidium and waterparks: an opportunity for disease prevention. Report to CDC, April. Majowicz, S.E., Michel, P., Aramini, J.J., McEwen, S.A. and Wilson, J.B. 2001. Descriptive analysis of endemic cryptosporidiosis cases reported in Ontario 1996–1997. Can. J. Pub. Health 92, 62–66. Mathieu, E., Levy, D.A., Veverka, F., Parrish, M.K., Sarisky, J., Shapiro, N., Johnston, S., Handzel, T., Hightower, A., Xiao, L., Lee, Y.M., York, S., Arrowood, M., Lee, R. and Jones, J.L. 2004. Epidemiologic and environmental investigation of a recreational water outbreak caused by two genotypes of Cryptosporidium parvum in Ohio in 2000. Am. J. Trop. Med. Hyg. 71, 582–589. McAnulty, J.M., Fleming, D.W. and Gonzalez, A.H. 1994. A community-wide outbreak of cryptosporidiosis associated with swimming at a wave pool. JAMA 272, 1597–1600. Mead, P.S., Slutsker, L., Dietz, V., McCaig, L.F., Bresee, J.S., Shapiro, C., Griffin, P.M. and Tauxe, R.V. 1999. Food-related illness and death in the United States. Emerg. Infect. Dis. 5, 607–625. Meisel, J.L., Perera, D.R., Meligro, C. and Rubin, C.E. 1976. Overwhelming watery diarrhea associated with a Cryptosporidium in an immunosuppressed patient. Gastroenterology 70, 1156–1160. 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. Moore, A.C., Herwaldt, B.L., Craun, G.F., Calderon, R.L., Highsmith, A.K. and Juranek, D.D. 1993. Surveillance for waterborne disease outbreaks—United States, 1991–1992. MMWR Surveill Summ. 42(No. SS5), 1–22. National Recreation and Park Association. 1999. The Encyclopedia of Aquatic Codes and Standards. NRPA, Ashburn, VA. Nazareth, B., Stanwell-Smith, R.E., Rowland, M.G. and O’Mahony, M.C. 1994. Surveillance of waterborne disease in England and Wales. Commun. Dis. Rep. CDR Rev. 4, R93–95.


366


Nelson, R., Norton, N., Cautley, E. and Furner, S. 1995. Community-based prevalence of anal incontinence. JAMA 274, 559–561. Nichols, G. 2006. Infection risks from water in natural and man-made environments. Euro. Surveill. 11(4), 76–78. Nichols, G., Chalmers, R., Lake, I., Sopwith, W., Regan, M., Hunter, P., Grenfell, P., Harrison, F. and Lane, C. 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. London, Drinking Water Inspectorate. Available at http://www.dwi.gov.uk/research/reports/DWI70_2_201.pdf. Nieminski, E.C. and Ongerth, J.E, 1995. Removing Giardia and Cryptosporidium by conventional treatment and direct filtration. J. AWWA 87, 96–106. 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. Okhuysen, P.C., Chappell, C.L., Crabb, J.H., Sterling, C.R. and DuPont, H.L. 1999. Virulence of three distinct Cryptosporidium parvum isolates for healthy adults. J. Infect. Dis. 180, 1275–1281. Oliveri, R., Di Piazza, F., Marsala, B., Cerame, G., Firenze, A. and Di Benedetto, M.A. 2006. Occurrence of Giardia cysts and Cryptosporidium oocysts in swimming pools in the province of Palermo, Italy. Ann. Ig. 18(5), 367–374. Ongerth, J.E. and Hutton, P.E. 1997. DE filtration to remove Cryptosporidium. J. AWWA 89, 39–46. Ongerth, J.E. and Hutton, P.E. 2001. Testing of diatomaceous earth filtration for removal of Cryptosporidium oocysts. J. AWWA 93, 54–63. Papapetropoulou, M. and Vantarakis, A.C. 1998 Detection of adenovirus outbreak at a municipal swimming pool by nested PCR amplification. J. Infect. 36, 101–103. Paterson, J. and Goldthorpe, I. 2006. Managing a cluster of cryptosporidiosis associated with a public swimming pool. NSW Pub. Health Bull. 17(5–6), 80. PHLS. 1995. Surveillance of waterborne diseases. Commun. Dis. Rep. CDR Wkly. 5(28), 129–130. PHLS. 1996a. Strength of association between human illness and water: revised definitions for use in outbreak investigations. Commun. Dis. Rep. CDR Wkly. 6(8), 65–67. PHLS. 1996b. Surveillance of waterborne disease and water quality. Commun. Dis. Rep. CDR Wkly. 6(8), 72–74. PHLS. 1997a. Surveillance of waterborne disease and water quality: July to December 1996. Commun. Dis. Rep. CDR Wkly. 7(8), 73–74. PHLS. 1997b. Surveillance of waterborne disease and water quality: January to June 1997. Commun. Dis. Rep. CDR Wkly. 7(35), 317–319. PHLS. 1998. Surveillance of waterborne disease and water quality: January to June 1998. Commun. Dis. Rep. CDR Wkly. 8(34), 305–306. PHLS. 1999a. Surveillance of waterborne disease and water quality: July to December 1998. Commun. Dis. Rep. CDR Wkly. 9(8), 73–75. PHLS. 1999b. Surveillance of waterborne disease and water quality: January to June 1999, and a summary of 1998. Commun. Dis. Rep. CDR Wkly. 9(34), 305–307. PHLS. 1999c. Cryptosporidiosis associated with swimming pools. Commun. Dis. Rep. CDR Wkly. 9(48), 423. PHLS. 2000a. Surveillance of waterborne disease and water quality: July to December 1999. Commun. Dis. Rep. CDR Wkly. 10(7), 65–67. PHLS. 2000b. British tourists return from Majorca with cryptosporidiosis. Commun. Dis. Rep. CDR Weekly 10(32), 285. PHLS. 2000c. Surveillance of waterborne disease and water quality: January to June 2000, and a summary of 1999. Commun. Dis. Rep. CDR Wkly. 10(35), 319–322. PHLS. 2001a. Surveillance of waterborne disease and water quality: July to December 2000. Commun. Dis. Rep. CDR Wkly. 11(7), 6–10. PHLS. 2001b. Surveillance of waterborne disease and water quality: January to June 2001, and a summary of 2000. Commun. Dis. Rep. CDR Wkly. 11(45), 5–10. PHLS. 2002a. Surveillance of waterborne disease and water quality: July to December 2001. Commun. Dis. Rep. CDR Wkly. 12(11), 11–16. PHLS. 2002b. Surveillance of waterborne disease and water quality: January to June 2002, and a summary of 2001. Commun. Dis. Rep. CDR Wkly. 12(37), 9–13.



367

PHLS. 2003. Surveillance of waterborne disease and water quality: January to June 2003, and a summary of 2002. Commun. Dis. Rep. CDR Wkly. 13(41), 10–14. PHLS. 2004. Surveillance of waterborne disease and water quality: July to December 2003. Commun. Dis. Rep. CDR Wkly. 14(15), 7–12. PHLS. 2006a. Surveillance of waterborne disease outbreaks summary of 2004. Commun. Dis. Rep. CDR Wkly. 16(15), 8–11. PHLS. 2006b. Surveillance of waterborne disease in England and Wales: summary of 2005. Commun. Dis. Rep. CDR Wkly. 16(28), 11–13. Pokorny, N.J., Weir, S.C., Carreno, R.A., Trevors, J.T. and Lee, H. 2002. Influence of temperature on Cryptosporidium parvum oocyst infectivity in river water samples as detected by tissue culture assay. J. Parasitol. 88, 641–643. Pool and Spa Marketing. 2003. United States swimming pool market. Hubbard Marketing and Publishing, Markham, Ontario, 20–21. Pool and Spa Marketing. 2006. Canadian swimming pool industry. Hubbard Marketing and Publishing, Markham, Ontario, 39–43. Pool Water Treatment Advisory Group. 1999. Swimming pool water, treatment and quality standards. Norfolk, Greenhouse Publishing. (www.pwtag.org./home.html). Pretlove, S.J., Radley, S., Toozs-Hobson, P.M., Thompson, P.J., Coomarasamy, A. and Khan, K.S. 2006. Prevalence of anal incontinence according to age and gender: a systematic review and meta-regression analysis. Int. Urogynecol. J. Pelvic Floor Dysfunct. 17, 407–417. Pruss, A. 1998. Review of epidemiological studies on health effects from exposure to recreational water. Int. J. Epidemiol. 27, 1–9. Puech, M.C., McAnulty, J.M., Lesjak, M., Shaw, N., Heron, L. and Watson, J.M. 2001. A statewide outbreak of cryptosporidiosis in New South Wales associated with swimming at public pools. Epidemiol. Infect. 126, 389–396. Robertson, B., Sinclair, M.I., Forbes, A.B., Veitch, M., Kirk, M., Cunliffe, D., Willis, J. and Fairley, C.K. 2002. Case-control studies of sporadic cryptosporidiosis in Melbourne and Adelaide, Australia. Epidemiol. Infect. 128, 419–431. Robertson, L.J., Campbell, A.T. and Smith, H.V. 1992. Survival of Cryptosporidium parvum oocysts under various environmental pressures. Appl. Environ. Microbiol. 58, 3494–3500. Rochelle, P.A., Upton, S.J., Montelone, B.A. and Woods, K. 2005. The response of Cryptosporidium parvum to UV light. Trends Parasitol. 21, 81–87. Rose, J.B. 1988. Occurrence and significance of Cryptosporidium in water. J. AWWA 80, 53–58. Rose, J.B., Lisle, J.T. and LeChevallier, M. 1997. Waterborne cryptosporidiosis: incidence, outbreaks, and treatment strategies. In Cryptosporidium and Cryptosporidiosis, ed. R. Fayer, 65–92. CRC Press, New York. Roy, S.L., DeLong, S.M., Stenzel, S., Shiferaw, B., Roberts, J., Khalakdina, A., Marcus, R., Nelson, R., Segler, S., Shah, D., Thomas, S., Vugia, D., Zansky, S., Dietz, V., Beach, M.J. and the Emerging Infections Program FoodNet Working Group. 2004. Risk factors for sporadic cryptosporidiosis among immunocompetent persons in the United States from 1999 to 2001. J. Clin. Microbiol. 42, 2944–2951. Roy, S.L., Scallan, E. and Beach, M.J. 2006. The rate of acute gastrointestinal illness in developed countries. J. Water Health. 4(Suppl 2), 31–69. Schaffzin, J.K., Keithly, J., Johnson, G., Sackett, D., Hoefer, D., Hoyt, L., Lurie, M., Teal, A., Rosen, B., Tavakoli, N., St. George, K., Braun-Howland, E. and Wallace, B. 2006. Large outbreak of cryptosporidiosis associated with a recreational water spraypark—New York, 2005. [abstract]. In Proceedings of the 55th Annual Epidemic Intelligence Service (EIS) Conference. Atlanta, Centers for Disease Control and Prevention. Schets, F.M., Engels, G.B. and Evers, E.G. 2004. Cryptosporidium and Giardia in swimming pools in the Netherlands. J. Water Health. 2, 191–200. Seyfried, P.L., Tobin, R.S., Brown, N.E. and Ness, P.F. 1985. A prospective study of swimming-related illness. I. Swimming-associated health risk. Am. J. Pub. Health. 75, 1068–1070. Smith, A., Reacher, M., Smerdon, W., Adak, G.K., Nichols, G. and Chalmers, R.M. 2006. Outbreaks of waterborne infectious intestinal disease in England and Wales, 1992–2003. Epidemiol. Infect. 134, 1141–1149.


368


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).



369

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.



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

371


372


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


Waste Management

373

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.


374 B.

Cryptosporidium and Cryptosporidiosis, Second Edition Effluent Treatment: Ozonation

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).


Waste Management E.

375

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).


376

IV.


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


Waste Management

377

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.


378 A.

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


Waste Management

379

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


380


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).


Waste Management

381

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


382


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.


Waste Management

383

References Adin, A. 2004. Particle count and size alteration for membrane fouling reduction in non-conventional water filtration. Wat. Sci. Technol. 50, 273–278. Atwill, E.R., Tate, K.W., Pereira, M.D.G.C., Bartolome, J. and Nader, G. 2006a. Efficacy of natural grasslandbuffers for removal of Cryptosporidium parvum in rangeland runoff. J. Food Protect. 69, 177–184. Atwill, E.R., Pereira, M.D.G.C., Alonso, L.H., Elmi, C., Epperson, W.B., Smith, R., Riggs, W., Carpenter, L.V., Dargatz, D.A. and Hoar, B. 2006b. Environmental load of Cryptosporidium parvum oocysts from cattle manure in feedlots from the central and western United States. J. Environ. Qual. 35, 200–206. Betancourt, W.Q. and Rose, J.B. 2004. Drinking water treatment processes for removal of Cryptosporidium and Giardia. Vet. Parasitol. 126, 219–234. Blackburn, B.G., Mazurek, J.M., Hlavsa, M., Park, J., Tillapaw, M., Parrish, M., Salehi, E., Franks, W., Koch, E., Smith, F., Xiao, L., Arrowood, M., Hill, V., da Silva, A., Johnston, S. and Jones, J.L. 2006. Cryptosporidiosis associated with ozonated apple cider. Emerg. Inf. Dis. 12, 684–686. Bonadonna, L., Briancesco, R., Ottaviani, M. and Veschetti, E. 2002. Occurrence of Cryptosporidium oocysts in sewage effluents and correlation with microbial, chemical, and physical water variables. Environ. Monitor. Assessment 75, 241–252. Bukhari, Z., Marshall, M.M., Korich, D.G., Fricker, C.R., Smith, H.V., Rosen, J. and Clancy, J.L. 2000. Comparison of Cryptosporidium parvum viability and infectivity assays following ozone treatment of oocysts. Appl. Environ. Microbiol. 66, 2972–2980. Bukhari, Z., Smith, H.V., Sykes, N., Humphreys, S.W., Paton, C.A., Girdwood, R.W.A. and Fricker, C.R. 1997. Occurrence of Cryptosporidium spp. oocysts and Giardia spp. cysts in sewage influents and effluents from treatment plants in England. Wat. Sci. Technol. 35, 385–390. Carraro, E., Fea, E., Salva, S. and Gili, G. 2000. Impact of a wastewater treatment plant on Cryptosporidium oocysts and Giardia cysts occurring in surface water. Wat. Sci. Technol. 41, 31–37. Charles, K., Ashbolt, N., Ferguson, C., Roser, D., McGuinness, R. and Deere, D. 2003. Centralised versus decentralised sewage systems: a comparison of pathogen and nutrient loads released into Sydney’s drinking water catchments. Wat. Sci. Technol. 48, 53–60. Chauret, C, Springthorpe, S. and Sattar, S. 1999. Fate of Cryptosporidium oocysts, Giardia cysts, and microbial indicators during wastewater treatment and anaerobic sludge digestion. Can. J. Microbiol. 45, 257–262. Collick A.S., Inglis, S., Steenhuis, T.S., Wright, P. and Bowman, D.D. (submitted) Inactivation of Ascaris suum in a biodrying compost system. J. Environ. Qual. Collick, A.S., Fogarty, E.A., Ziegler, P.E., Walter, M.T., Bowman, D.D. and Steenhuis, T.S. 2006. Survival of Cryptosporidium parvum oocysts in calf housing facilities in the New York City watersheds. J. Environ. Qual. 35, 680–687. Corona-Vasquez, B., Samuelson, A., Rennecker, J.L. and Marinas, B.J. 2002. Inactivation of Cryptosporidium parvum oocysts with ozone and free chlorine. Wat. Res. 36, 4053–4063. Côté, C., Massé, D.I. and Quessy, S. 2006a. Reduction of indicator and pathogenic microorganisms by psychrophilic anaerobic digestion in swine slurries. Biores. Technol. 97, 686–691. Côté, C., Villeneuve, A., Lessard, L. and Quessy, S. 2006b. Fate of pathogenic and nonpathogenic microorganisms during storage of liquid hog manure in Québec. Livestock Sci. 102, 204–210. Dai, X. and Boll, J. 2006. Settling velocity of Cryptosporidium parvum and Giardia lamblia. Wat. Res. 40, 1321–1325. Fayer, R. 2007. The general biology of Cryptosporidium. In Cryptosporidium and cryptosporidiosis, 2nd Ed., Fayer, R.and Xiao, L. eds. CRC Press, Boca Raton, FL. Fayer, R., Santín, M, Trout, J.M. and Greiner, E. 2006. Prevalence of species and genotypes of Cryptosporidium found in 1–2-year-old dairy cattle in the eastern United States. Vet. Parasitol. 135, 105–112. Fayer, R., Santín, M. and Xiao, L. 2005. Cryptosporidium bovis n. sp. (Apicomplexa: Cryptosporidiidae) in cattle (Bos taurus). J. Parasitol. 91, 624–629. Fayer, R., Trout, J.M., Walsh, E. and Cole, R. 2000. Rotifers ingest oocysts of Cryptosporidium parvum. J. Eukaryot. Microbiol. 47, 161–163. Fox, J.C., Fitzgerald, P.R. and Lue-Hing, C. 1981. Sewage Organisms: A Color Atlas. The Metropolitan Sanitary District of Greater Chicago, Chicago, IL, 116 pp.


384


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.


Waste Management

385

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.



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

387


388


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


Fish, Amphibians, and Reptiles

389

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


390


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).


Fish, Amphibians, and Reptiles 6.

391

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.


392 E.

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.


Fish, Amphibians, and Reptiles

393

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.


394


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.


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.

395


396


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,


Birds

397

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


398


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


Birds

399

(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 60%) from 1000 pseudoreplicates is indicated at the left of the supported node.


Birds

405

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,


406


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).


Birds

407

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


408


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


Birds

409

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


410


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.


Birds

411

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


412


(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


Birds

413

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.

References Abbassi, H., Coudert, F., Cherel, Y., Dambrine, G., Brugere-Picoux, J. and Naciri, M. 1999. Renal cryptosporidiosis (Cryptosporidium baileyi) in specific-pathogen-free chickens experimentally coinfected, with Marek’s disease virus. Avian Dis. 43, 738–744. Abbassi, H., Dambrine, G., Cherel, Y., Coudert, F. and Naciri, M. 2000. Interaction of Marek’s disease virus and Cryptosporidium baileyi in experimentally infected chickens. Avian Dis. 44, 776–789. Abe, N. and Iseki, M. 2004. Identification of Cryptosporidium isolates from cockatiels by direct sequencing of the PCR-amplified small subunit ribosomal RNA gene. Parasitol. Res. 92, 523–526. 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. Belton, D.J. and Powell, I.B. 1987. Cryptosporidiosis in Lovebirds (Agapornis sp.). N. Z. Vet. J. 35, 15–15. Bermudez, A.J., Ley, D.H., Levy, M.G., Ficken, M.D., Guy, J.S. and Gerig, T.M. 1988. Intestinal and bursal Cryptosporidiosis in turkeys following inoculation with Cryptosporidium sp. isolated from commercial poults. Avian Dis. 32, 445–450. Blagburn, B.L., Lindsay, D.S., Giambrone, J.J., Sundermann, C.A. and Hoerr, F.J. 1987. Experimental cryptosporidiosis in broiler-chickens. Poult. Sci. 66, 442–449. Blagburn, B.L., Lindsay, D.S., Hoerr, F.J., Atlas, A.L. and Toiviokinnucan, M. 1990. Cryptosporidium sp infection in the proventriculus of an Australian diamond firetail finch (Staganoplura bella Passeriformes, Estrildidae). Avian Dis. 34, 1027–1030. Cama, V.A., Bern, C., Sulaiman, I.M., Gilman, R.H., Ticona, E., Vivar, A., Kawai, V., Vargas, D., Zhou, L. and Xiao, L.H. 2003. Cryptosporidium species and genotypes in HIV-positive patients in Lima, Peru. J. Eukaryot. Microbiol. 50, 531–533. Cardozo, S.V., Teixeira Filho, W.L. and Lopes, C.W. 2005. [Experimental transmission of Cryptosporidium baileyi (Apicomplexa: Cryptosporidiidae) isolated of broiler chicken to Japanese quail (Coturnix japonica)]. Rev. Bras. Parasitol. Vet. 14, 119–124. Champliaud, D., Gobet, P., Naciri, M., Vagner, O., Lopez, J., Buisson, J.C., Varga, I., Harly, G., Mancassola, R. and Bonnin, A. 1998. Failure to differentiate Cryptosporidium parvum from C. meleagridis based on PCR amplification of eight DNA sequences. Appl. Environ. Microbiol. 64, 1454–1458. Cheadle, M.A., Toivio-Kinnucan, M. and Blagburn, B.L. 1999. The ultrastructure of gametogenesis of Cryptosporidium baileyi (Eimeriorina; Cryptosporidiidae) in the respiratory tract of broiler chickens (Gallus domesticus). J. Parasitol. 85, 609–615. Chvala, S., Fragner, K., Hackl, R., Hess, M. and Weissenbock, H. 2006. Cryptosporidium infection in domestic geese (Anser anser f. domestica) detected by in situ hybridization. J. Comp. Pathol. 134, 211–218. Coupe, S., Sarfati, C., Hamane, S. and Derouin, F. 2005. Detection of Cryptosporidium and identification to the species level by nested PCR and restriction fragment length polymorphism. J. Clin. Microbiol. 43, 1017–1023. 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. Current, W.L. and Snyder, D.B. 1988. Development of and serologic evaluation of acquired immunity to Cryptosporidium baileyi by broiler chickens. Poult. Sci. 67, 720–729. Darabus, G. and Olariu, R. 2003. The homologous and interspecies transmission of Cryptosporidium parvum and Cryptosporidium meleagridis. Pol. J. Vet. Sci. 6, 225–228.


414


Dhillon, A.S., Thacker, H.L., Dietzel, A.V. and Winterfield, R.W. 1981. Respiratory cryptosporidiosis in broiler chickens. Avian Dis. 25, 747–751. Egyed, Z., Sreter, T., Szell, Z., Beszteri, B., Dobos-Kovacs, M., Marialigeti, K., Cornelissen, A.W.C.A. and Varga, I. 2002. Polyphasic typing of Cryptosporidium baileyi: A suggested model for characterization of cryptosporidia. J. Parasitol. 88, 237–243. Enemark, H.L., Ahrens, P., Juel, C.D., Petersen, E., Petersen, R.F., Andersen, J.S., Lind, P. and Thamsborg, S.M. 2002. Molecular characterization of Danish Cryptosporidium parvum isolates. Parasitology 125, 331–341. Fayer, R., Speer, C. and Dubey, J., 1997. The general biology of Cryptosporidium in Cryptosporidium and Cryptosporidiosis, Fayer, R., Eds. CRC Press, Boca Raton, FL, pp. 1–41. Fernandez, A., Quezada, M., Gomez, M.A., Navarro, J.A., Rodriguez, J. and Sierra, M.A. 1990. Cryptosporidiosis in chickens from southern Spain. Avian Dis. 34, 224–227. Gajadhar, A.A. 1994. Host specificity studies and oocyst description of a Cryptosporidium sp isolated from ostriches. Parasitol. Res. 80, 316–319. Gardiner, C.H. and Imes, G.D., Jr. 1984. Cryptosporidium sp. in the kidneys of a black throated finch. J. Am. Vet. Med. Assoc. 185, 1401–1402. Gatei, W., Suputtamongkol, Y., Waywa, D., Ashford, R.W., Bailey, J.W., Greensill, J., Beeching, N.J. and Hart, C.A. 2002. Zoonotic species of Cryptosporidium are as prevalent as the anthroponotic in HIV-infected patients in Thailand. Ann. Trop. Med. Parasitol. 96, 797–802. Gatei, W., Greensill, J., Ashford, R.W., Cuevas, L.E., Parry, C.M., Cunliffe, N.A., Beeching, N.J. and Hart, C.A. 2003. Molecular analysis of the 18S rRNA gene of Cryptosporidium parasites from patients with or without human immunodeficiency virus infections living in Kenya, Malawi, Brazil, the United Kingdom, and Vietnam. J. Clin. Microbiol. 41, 1458–1462. Gatei, W., Das, P., Dutta, P., Sen, A., Cama, V., Lal, A.A. and Xiao, L. 2006a. Multilocus sequence typing and genetic structure of Cryptosporidium hominis from children in Kolkata, India. Infect. Genet. Evol. 7, 197–205. Gatei, W., Wamae, C.N., Mbae, C., Waruru, A., Mulinge, E., Waithera, T., Gatika, S.M., Kamwati, S.K., Revathi, G. and Hart, C.A. 2006b. Cryptosporidiosis: Prevalence, genotype analysis, and symptoms associated with infections in children in Kenya. Am. J. Trop. Med. Hyg. 75, 78–82. Gharagozlou, M. and Khodashenas, M. 1985. Cryptosporidiosis in a native rooster with a chronic proliferative enteritis. Arch. Vet. 17, 129. Gharagozlou, M.J., Dezfoulian, O., Rahbari, S., Bokaie, S., Jahanzad, I. and Razavi, A.N. 2006. Intestinal cryptosporidiosis in turkeys in Iran. J. Vet. Med. A Physiol. Pathol. Clin. Med. 53, 282–285. Glaberman, S., Sulaiman, I.M., Bern, C., Limor, J., Peng, M.M., Morgan, U., Gilman, R., Lal, A.A. and Xiao, L.H. 2001. A multilocus genotypic analysis of Cryptosporidium meleagridis. J. Eukaryot. Microbiol., 19s–22s. Glisson, J.R., Brown, T.P., Brugh, M., Page, R.K., Kleven, S.H. and Davis, R.B. 1984. Sinusitis in turkeys associated with respiratory cryptosporidiosis. Avian Dis. 28, 783–790. Goodwin, M.A., Latimer, K.S., Brown, J., Steffens, W.L., Martin, P.W., Resurreccion, R.S., Smeltzer, M.A. and Dickson, T.G. 1988a. Respiratory cryptosporidiosis in chickens. Poult. Sci. 67, 1684–1693. Goodwin, M.A., Steffens, W.L., Russell, I.D. and Brown, J. 1988b. Diarrhea associated with intestinal cryptosporidiosis in turkeys. Avian Dis. 32, 63–67. Goodwin, M.A. and Krabill, V.A. 1989. Diarrhea associated with small intestinal cryptosporidiosis in a budgerigar and in a cockatiel. Avian Dis. 33, 829–833. Graczyk, T.K., Cranfield, M.R., Fayer, R., Trout, J. and Goodale, H.J. 1997. Infectivity of Cryptosporidium parvum oocysts is retained upon intestinal passage through a migratory water-fowl species (Canada goose, Branta canadensis). Trop. Med. Int. Health 2, 341–347. Guy, J.S., Levy, M.G., Ley, D.H., Barnes, H.J. and Gerig, T.M. 1987. Experimental reproduction of enteritis in bobwhite quail (Colinus virginianus) with Cryptosporidium and reovirus. Avian Dis. 31, 713–722. Guyot, K., Follet-Dumoulin, A., Lelievre, E., Sarfati, C., Rabodonirina, M., Nevez, G., Cailliez, J.C., Camus, D. and Dei-Cas, E. 2001. Molecular characterization of Cryptosporidium isolates obtained from humans in France. J. Clin. Microbiol. 39, 3472–3480. Hajdusek, O., Ditrich, O. and Slapeta, J. 2004. Molecular identification of Cryptosporidium spp. in animal and human hosts from the Czech Republic Vet. Parasitol. 122, 183–192.


Birds

415

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.


416


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.


Birds

417

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.


418


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.


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


420


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


Zoo and Wild Mammals

421

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


422


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,



423

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


424


(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,



425

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).


426


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).



427

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,


428


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



429

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


430


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.

References Abe, N. and Iseki, M. 2003. Identification of genotypes of Cryptosporidium parvum isolates from ferrets in Japan. Parasitol. Res. 89, 422–424. Abe, N., Matsubayashi, M., Kimata, I. and Iseki, M. 2006. Subgenotype analysis of Cryptosporidium parvum isolates from humans and animals in Japan using the 60-kDa glycoprotein gene sequences. Parasitol. Res. 99, 303–305. Abe, N., Takami, K., Kimata, I. and Iseki, M. 2004. Molecular characterization of a Cryptosporidium isolate from a banded mongoose Mungos mungo. J. Parasitol. 90, 167–171. Alves, M., Matos, O., Fonseca, I.P., Delgado, E., Lourenço, A.M. and Antunes, F. 2001. Multilocus genotyping of Cryptosporidium isolates from human HIV-infected and animal hosts. J. Euk. Microbiol. (Suppl.), 17 and 18S. Alves, M., Xiao, L., Antunes, F. and Matos, O. 2006. Distribution of Cryptosporidium subtypes in humans and domestic and wild ruminants in Portugal. Parasitol. Res. 99, 287–292. Alves, M., Xiao, L., Lemos, V., Zhou, L., Cama, V., da Cunha, MB., Matos, O. and Antunes, F. 2005. Occurrence and molecular characterization of Cryptosporidium spp. in mammals and reptiles at the Lisbon Zoo. Parasitol. Res. 97, 108–112. Alves, M., Xiao, L., Sulaiman, I., Lal, A.A., Matos, O. and Antunes, F. 2003. Subgenotype analysis of Cryptosporidium isolates from humans, cattle, and zoo ruminants in Portugal. J. Clin. Microbiol. 41, 2744–2747. Anderson, B.C. 1987. Abomasal cryptosporidiosis in cattle. Vet. Pathol. 24, 235–238. Appelbee, A.J., Thompson, R.C. and Olson, M.E. 2005. Giardia and Cryptosporidium in mammalian wildlife—current status and future needs. Trends Parasitol. 21, 370–376. Atwill, E.R., Phillips, R., Pereira, M.D., Li, X. and McCowan, B. 2004. Seasonal shedding of multiple Cryptosporidium genotypes in California ground squirrels (Spermophilus beecheyi). Appl. Environ. Microbiol. 70, 6748–6752.



431

Atwill, E.R., Camargo, S.M., Phillips, R., Alonso, L.H., Tate, K.W., Jensen, W.A., Bennet, J., Little, S. and Salmon, T.P. 2001. Quantitative shedding of two genotypes of Cryptosporidium parvum in California ground squirrels (Spermophilus beecheyi). Appl. Environ. Microbiol. 67, 2840–2843. Atwill, E.R., Sweitzer, R.A., Pereira, M.G., Gardner, I.A., Van Vuren, D. and Boyce, W.M. 1997. Prevalence of associated risk factors for shedding Cryptosporidium parvum oocysts and Giardia cysts within feral pig populations in California. Appl. Environ. Microbiol. 63, 3946–3949. Bajer, A., Bednarska, M., Pawelczyk, A., Behnke, J.M., Gilbertn, F.S. and Sinski, E. 2002. Prevalence and abundance of Cryptosporidium parvum and Giardia spp. in wild rural rodents from the Mazury Lake District region of Poland. Parasitology. 125, 21–34. Bajer, A., Cacciò, S., Bednarska, M., Behnke, J.M., Pieniazek, N.J. and Sinski, E. 2003. Preliminary molecular characterization of Cryptosporidium parvum isolates of wildlife rodents from Poland. J. Parasitol. 89, 1053–1055. Barker, I.K., Beveridge, I., Bradley, A.J. and Lee, A.K. 1978. Observations on spontaneous stress-related mortality among males of the dasyurid marsupial, Antechinus stuartii Macleay. Aust. J. Zool. 26, 435. Bednarska, M., Bajer, A., Kulis, K. and Sinski, E. 2003. Biological characterisation of Cryptosporidium parvum isolates of wildlife rodents in Poland. Ann. Agric. Environ. Med. 10, 163–169. Bertolino, S., Wauters, L.A., De Bruyn, L. and Canestri-Trotti, G. 2003. Prevalence of coccidia parasites (Protozoa) in red squirrels (Sciurus vulgaris): effects of host phenotype and environmental factors. Oecologia 137, 286–295. Bull, S.A., Chalmers, R.M., Sturdee, A.P. and Healing, T.D. 1998. A survey of Cryptosporidium species in Skomer bank voles Clethrionomys glareolus skomerensis. J. Zool. Lond. 244, 119–122. Carlson, B.L. and Nielsen, S.W. 1982. Cryptosporidiosis in a raccoon. J. Am. Vet. Med. Assoc. 181, 1405–1406. Casemore, D.P., Wright, S.E. and Coop, R.L. 1997. Cryptosporidiosis—human and animal epidemiology, in Cryptosporidium and Cryptosporidiosis, Fayer, R., ed., CRC Press, Boca Raton, Florida, 65–92. Chalmers, R.M., Sturdee, A.P., Bull, S.A., Miller, A. and Wright, S.E. 1997. The prevalence of Cryptosporidium parvum and C. muris in Mus domesticus, Apodemus sylvaticus, and Clethrionomys glareolus in an agricultural system. Parasitol. 83, 478–482. Chalmers, R.M., Sturdee, A.P., Bull, S.A. and Miller, A.M., 1995. Rodent reservoirs of Cryptosporidium, in Protozoan Parasites and Water, Betts, W.B., Casemore, D., Fricker, C., Smith, H., Watkins, J., eds., The Royal Society of Chemistry Special Publication, No. 168, CUP, UK. 63–66. Chalmers, R.M., Sturdee, A.P., Casemore, D.P., Curry, A., Miller, A., Parker, N.D. and Richmond, T.M. 1994. Cryptosporidium muris in wild house mice (Mus musculus) First report in the U.K. Eur. J. Protistol. 30. 151–155. Chilvers, B.L., Cowan, P.E., Waddington, D.C., Kelly, P.J. and Brown, T.J. 1998. The prevalence of infection of Giardia spp. and Cryptosporidium spp. in wild animals on farmland, southeastern North Island, New Zealand. Int. J. Environ. Health Res. 8, 59–64. Cox, P., Griffith, M., Angles, M., Deere, D. and Ferguson, C. 2005. Concentrations of pathogens and indicators in animal feces in the Sydney watershed. Appl. Environ. Microbiol. 71, 5929–5934. Da Silva, A.J., Caccio, S., Williams, C., Won, K.Y., Nace, E.K., Whittier, C., Pieniazek, N.J. and Eberhard, M.L. 2003. Molecular and morphologic characterization of a Cryptosporidium genotype identified in lemurs. Vet. Parasitol. 111, 297–307. DaSilva, N.R.S., Bigatti, L.E., Amado, R.K., DeAraujo, F.A.P. and Chaplin, E.L. 1992. Eimeria and Cryptosporidium infections in rabbits in the municipality of Porto Alegre, Rio Grande, Brazil. Arq. Faculd. Vet. UFRGS 20, 235. Delgado, E., Fonseca, I.P., Fazendeiro, I., Matos, O., Antunes, F. and Cunha, M.B. 2003 Cryptosporidium spp. in ruminants at the Lisbon Zoo. J. Zoo Wild. Med. 34, 352–356. Deng, M.Q. and Cliver, D.O. 1999. Improved immunofluorescence assay for detection of Giardia and Cryptosporidium from asymptomatic adult cervine animals. Parasitol. Res. 85, 733–736. Deng, M.Q., Lam, K.M. and Cliver, D.O. 2000. Immunomagnetic separation of Cryptosporidium parvum oocysts using MACS MicroBeads and high gradient separation columns. J. Microbiol. Methods. 40, 11–17. Dubey, J.P., Hamir, A.N., Sonn, R.J. and Topper, M. J. 1998. Cryptosporidiosis in a bat (Eptesicus fuscus). J. Parasitol. 84, 622 and 623. Dubey, J.P., Markovits, J.E. and Killary, K.A. 2002. Cryptosporidium muris-like in stomach of cynomolgus monkeys (Macaca fascicularis). Vet. Pathol. 39, 363–371.


432


Ekanayake, D.K., Arulkanthan, A., Horadagoda, N.U., Sanjeevani, G.K., Kieft, R., Gunatilake, S. and Dittus, W.P. 2006. Prevalence of Cryptosporidium and other enteric parasites among wild nonhuman primates in Polonnaruwa, Sri Lanka. Am. J. Trop. Med. Hyg. 74, 322–329. Estes, R.D., Otte, D. and Wilson, E. 1992. The Behaviour Guide to African Mammals, including Hoofed Mammals, Carnivores and Primates. 1st ed. The University of California Press, California, USA. 660. Fayer, R. 2004. Cryptosporidium: a waterborne zoonotic parasite. Vet. Parasitol. 126, 37–56. Fayer, R. and Ungar B.L.P. 1986. Cryptosporidium spp. and cryptosporidiosis. Microbiol. Rev. 50, 458–483. Fayer, R., Fisher, J.R., Sewell, C.T., Kavanaugh, D.M. and Osborn, D.A. 1996. Spontaneous cryptosporidiosis in captive white-tailed deer (Odocoileus virginianus). J. Wildl. Dis. 32, 619–622. Fayer, R., Phillips, L., Anderson, B.C. and Bush, M. 1991. Chronic cryptosporidiosis in a Bactrian camel (Camelus bactrianus). J. Zoo Wildl. Med. 22, 228–232. Fayer, R., Santín, M. and Xiao, L. 2005. Cryptosporidium bovis n. sp. (Apicomplexa: Cryptosporidiidae) in cattle (Bos taurus). J. Parasitol. 91, 624–629. Fayer, R., Santín, M., Trout, J.M., DeStefano, S., Koenen, K., Kaur, T. 2006. Prevalence of Microsporidia, Cryptosporidium spp., and Giardia spp. in beavers (Castor canadensis) in Massachussetts. J. Zoo and Wildlife Med. 37, 492–497. 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, Florida. 1–41. Gatei, W., Greensill, J., Ashford, R.W., Cuevas, L.E., Parry, C.M., Cunliffe, N.A., Beeching, N.J. and Hart, C.A. 2003. Molecular analysis of the 18S rRNA gene of Cryptosporidium parasites from patients with or without human immunodeficiency virus infections living in Kenya, Malawi, Brazil, the United Kingdom, and Vietnam, J. Clin. Microbiol. 41, 1458–1462. Gómez, M.S., Gracenea, M., Gosalbez, P., Feliu, C., Enseñat, C. and Hidalgo, R. 1992. Detection of oocysts of Cryptosporidium in several species of monkeys and in one prosimian species at the Barcelona Zoo. Parasitol. Res. 78, 619 and 620. Gómez, M.S., Torres, J., Gracenea, M., Fernandez-Morán, J. and Gonzalez-Moreno, O. 2000. Further report on Cryptosporidium in Barcelona Zoo. Parasitol. Res. 86, 318–323. Gómez, M.S., Vila, T., Feliu, C., Montoliu, I., Gracenea, M. and Fernandez, F. 1996. A survey for Cryptosporidium spp. in mammals at the Barcelona Zoo. Int. J. Parasitol. 26, 1331–1333. Gracenea, M., Gomez, M.S., Torres, J., Carne, E. and Fernandez-Moran, J. 2002. Transmission dynamics of Cryptosporidium in primates and herbivores at the Barcelona zoo: a long-term study. Vet. Parasitol. 104, 19-26. Graczyk, T.K., Cranfield, M.R., Dunning, C. and Strandberg, J.D. 1998. Fatal cryptosporidiosis in a juvenile captive African hedgehog (Ateletrix albiventris). J. Parasitol. 84, 178–180. Graczyk, T.K., Da Silva, A.J., Cranfield, M.R., Nizeyi, J.B., Kalema, G.R. and Pieniazek, N.J. 2001. Cryptosporidium parvum genotype 2 infections in free-ranging mountain gorillas (Gorilla gorilla beringei) of the Bwindi Impenetrable National Park, Uganda. Parasitol. Res. 87, 368–370. Hajdusek, O., Ditrich, O. and Slapeta, J. 2004. Molecular identification of Cryptosporidium spp. in animal and human hosts from the Czech Republic. Vet. Parasitol. 122, 183–192. Hamnes, I.S., Gjerde, B., Robertson, L., Vikoren, T. and Handeland, K. 2006. Prevalence of Cryptosporidium and Giardia in free-ranging wild cervids in Norway. Vet. Parasitol. 141, 30–41. Hampton, J.C. and Rosário, B. 1966. The attachment of protozoan parasites to intestinalepithelial cells of the mouse. J. Parasitol. 52, 939–949. Harp, J.A. and Goff, J. P. 1998. Strategies for the control of Cryptosporidium parvum infections in calves. J. Dairy Sci. 81, 289–294. Heitman, T.L., Frederick, L.M., Viste, J.R., Guselle, N.J., Morgan, U.M., Thompson, R. C.A., and Olson, M.E. 2002. Prevalence of Giardia and Cryptosporidium and characterization of Cryptosporidium spp. isolated from wildlife, human, and agricultural sources in the North Saskatchewan River Basin in Alberta, Canada. Can. J. Microbiol. 48, 530–541. Heuschele, W.P., Oosterhuis, J., Janssen, D., Robinson, P.T., Ensley, P.K., Meier E., Olson, T., Anderson, M.P. and Benirschke, K. 1986. Cryptosporidial infections in captive wild animals. J. Wildl. Dis. 22, 493–496. Hikosaka, K. and Nakai, Y. 2005. A novel genotype of Cryptosporidium muris from large Japanese field mice, Apodemus speciosus. Parasitol. Res. 97, 373–379. Hope, K., Goldsmith, M.L. and Graczyk, T. 2004. Parasitic health of olive baboons in Bwindi Impenetrable National Park, Uganda. Vet. Parasitol.122, 165–170.



433

ˇ ´ D. 2003. Natural infection of Cryptosporidium muris (Apicomplexa: Hurková, L., Hajdusek, O. and Modry, Cryptosporiidae) in Siberian chipmunks. J. Wildl. Dis. 39, 441–444. Inman, L.R. and Takeuchi A. 1979. Spontaneous cryptosporidiosis in an adult female rabbit. Vet. Pathol. 16, 89–95. Iseki, M. 1986. Two species of Cryptosporidium naturally infecting house rats, Rattus-norvegicus. Jpn. J. Parasitol. 35, 521–526. Iseki, M., Maekawa, T., Mriya, K., Uni, S. and Takada, S. 1989. Infectivity of Cryptosporidium muris (strain RN 66) in various laboratory animals. Parasitol. Res. 75, 218–222. Jellison, K.L., Hemond, H.F. and Schauer, D.B. 2002. Sources and species of Cryptosporidium oocysts in the Wachusett Reservoir watershed. Appl. Environ. Microbiol. 68, 569–575. Jiang, J., Alderisio, K.A. and Xiao, L. 2005. Distribution of Cryptosporidium genotypes in storm-event water samples from three watersheds in New York. Appl. Environ. Microbiol. 71, 4446–4454. Juranek, D.D. 1995. Cryptosporidiosis: sources of infection and guidelines for prevention. Clin. Infect. Dis. 21, 557–561. Kalishman, J., Paul-Murphy, J., Scheffler, J. and Thomson, J.A. 1996. Survey of Cryptosporidium and Giardia spp. in a captive population of common marmosets. Lab. Anim. Sci. 46, 116–119. Katsumata, T., Hosea, D., Ranuh, I.G., Uga, S., Yanagi, T. and Kohno, S. 2000. Short report: possible Cryptosporidium muris infection in humans. Am. J. Trop. Med. Hyg. 62, 70–72. Klesius, P.H., Haynes, T.B. and Malo, L.K. 1986. Infectivity of Cryptosporidium sp. isolated from wild mice for calves and mice. J. Am. Vet. Med. Assoc. 189, 192 and 193. Korsholm, H. and Henriksen, S.A. 1984. Infection with Cryptosporidium in roe deer (Capreolus capreolus L.). A preliminary report. Nord. Vet. Med. 36, 266. Kovatch, R.M. and White, J.D. 1972. Cryptosporidiosis in two juvenile rhesus monkeys. Veterinary Pathology 9, 426–440. Kruse, H., Kirkemo, A.M. and Handeland, K. 2004. Wildlife as source of zoonotic infections. Emerg. Infect. Dis. 10, 2067–2072. Laakkonen, J., Soveri, T. and Hentotten, H. 1994. Prevalence of Cryptosporidium sp. in peak density Microtus agrestis, M. oeconamus, and Clethrionomys glareolus populations. J. Wildl. Dis. 30, 110 and 111. Legesse, M. and Erko, B. 2004. Zoonotic intestinal parasites in Papio anubis (baboon) and Cercopithecus aethiops (vervet) from four localities in Ethiopia. Acta Trop. 90, 231–236. Levine, N. D. 1980. Some corrections of coccidian (Apicomplexa: Protozoa) nomenclature. J. Parasitol. 66, 830–834. Lindsay, D.S., Hendrix, C.M. and Blagburn, B.L. 1988. Experimental Cryptosporidium parvum infections in opossums (Didelphis virginiana). J. Wildl. Dis. 24, 157–159. Lourenço, A.M., Bruno de Sousa, C, Matos, O., Alves, M., Pereira da Fonseca, I. and Antunes, F. 2000. Estudo preliminar da criptosporidiose em gamos (Dama dama L.) da Tapada Nacional de Mafra (Portugal). Acta Parasitol. Port. 7, 29–31. McGuigan, C. 2005. Cryptosporidium outbreak after a visit to a wildlife centre in northeast Scotland: 62 confirmed cases. Euro. Surveill. 2005. 10, E050428.2. Majewska, A.C., Kasprzak, W. and Werner, A. 1997. Prevalence of Cryptosporidium in mammals housed in Poznan´ Zoological Garden, Poland. Acta Parasitol. 42, 195–198. Martin, H.D. and Zeidner, N.S. 1992. Concomitant cryptosporidia, coronavirus, and parvovirus infection in a raccoon (Procyon lotor). J. Wildl. Dis. 28, 113–115. Matsubayashi, M., Abe, N., Takami, K., Kimata, I., Iseki, M., Nakanishi, T., Tani, H., Sasai, K. and Baba, E. 2004 . First record of Cryptosporidium infection in a raccoon dog (Nyctereutes procyonoides viverrinus). Vet. Parasitol. 120, 171–175. Matsui, T., Fujino, T., Kajima, J. and Tsuji, M. 2000. Infectivity to experimental rodents of Cryptosporidium parvum oocysts from Siberian chipmunks (Tamias sibiricus) originated in the People’s Republic of China. J. Vet. Med. Sci. 62, 487–489. Miller, R.A., Bronsdon, M.A., Kuller, L. and Morton, W.R. 1990. Clinical and parasitologic aspects of cryptosporidiosis in nonhuman primates. Science. 40, 42–46. Miller, R.E. 1999. Quarantine: a necessity for zoo and aquarium animals, in Zoo and Wild Animal Medicine, Fowler, M.E. and Miller, R.E., Eds., W.B. Saunders, Philadelphia, Pennsylvania. 57–65. Miyaji, S., Tanikawa, T. and Shikata, J. 1989. Prevalence of Cryptosporidium in Rattus rattus and R. norvegicus in Japan. Jpn. J. Parasitol. 38, 368–372.


434


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.



435

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.


436


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.

52268_C017.fm Page 437 Tuesday, September 4, 2007 7:41 AM

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


438

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


Companion Animals

439

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.


440


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


Companion Animals

441

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 (

Cryptosporidium and Cryptosporidiosis

Giardia and Cryptosporidium from Molecules to Disease

Cryptosporidium: The Analytical Challenge (Special Publication)

The Pathogenic Enteric Protozoa:: Giardia, Entamoeba, Cryptosporidium and Cyclospora (World Class Parasites)

The Pathogenic Enteric Protozoa:: Giardia, Entamoeba, Cryptosporidium and Cyclospora (World Class Parasites)

Pride And Prejudice And Zombies

Pride and Prejudice and Zombies

Pride And Prejudice And Zombies

Lions And Tigers And Snares

Pride and Prejudice and Zombies

Pride and Prejudice and Zombies

Pride and Prejudice and Zombie

Pride And Prejudice And Zombies

Pride and Prejudice and Zombies

Theogony and Works and Days

Pride And Prejudice And Zombies

PRIDE AND PREJUDICE AND ZOMBIES

Humans and Demons and Elves

Pride and Prejudice and Zombies

Pride and Prejudice and Zombies

Pride and Prejudice and Zombies

Pride And Prejudice And Zombies

Consciousness and Persons: Unity and Identity (Representation and Mind)

Milk Consumption and Health (Food and Beverage Consumption and Health)

Abuse, Torture and Trauma, and Their Consequences and Effects

Man and the Beasts (Medieval and Renaissance Texts and Studies)

Environmental and Health Risk Assessment and Management: Principles and Practices

The Media and Hurricanes Katrina and Rita: Lost and Found

Normal and Abnormal Fear and Anxiety in Children and Adolescents

Cryptosporidium and Cryptosporidiosis

Giardia and Cryptosporidium from Molecules to Disease

Cryptosporidium: The Analytical Challenge (Special Publication)

The Pathogenic Enteric Protozoa:: Giardia, Entamoeba, Cryptosporidium and Cyclospora (World Class Parasites)

The Pathogenic Enteric Protozoa:: Giardia, Entamoeba, Cryptosporidium and Cyclospora (World Class Parasites)

Pride And Prejudice And Zombies

Pride and Prejudice and Zombies

Pride And Prejudice And Zombies

Lions And Tigers And Snares

Pride and Prejudice and Zombies

Pride and Prejudice and Zombies

Pride and Prejudice and Zombie

Pride And Prejudice And Zombies

Pride and Prejudice and Zombies

Theogony and Works and Days

Pride And Prejudice And Zombies

PRIDE AND PREJUDICE AND ZOMBIES

Humans and Demons and Elves

Pride and Prejudice and Zombies

Pride and Prejudice and Zombies

Pride and Prejudice and Zombies

Pride And Prejudice And Zombies

Consciousness and Persons: Unity and Identity (Representation and Mind)

Milk Consumption and Health (Food and Beverage Consumption and Health)

Abuse, Torture and Trauma, and Their Consequences and Effects

Man and the Beasts (Medieval and Renaissance Texts and Studies)

Environmental and Health Risk Assessment and Management: Principles and Practices

The Media and Hurricanes Katrina and Rita: Lost and Found

Normal and Abnormal Fear and Anxiety in Children and Adolescents

Recommend Documents