Giardia and Cryptosporidium from Molecules to Disease

GIARDIA AND CRYPTOSPORIDIUM From Molecules to Disease

Acknowledgement We are most grateful to Arturo Pérez-Taylor for his excellent informatics assistance during the preparation of this book.

GIARDIA AND CRYPTOSPORIDIUM From Molecules to Disease

Edited by

Guadalupe Ortega-Pierres Department of Genetics and Molecular Biology, CINVESTAV-IPN, México City, Mexico

Simone M. Cacciò Department of Infectious, Parasitic and Immunomediated Diseases, Istituto Superiore di Sanità, Rome, Italy

Ronald Fayer Environmental Microbial Safety Laboratory, Animal and Natural Resources Institute, ARS-USDA, Beltsville, MD, USA

Theo G. Mank Department of Parasitology, Public Health Laboratory, Haarlem, the Netherlands

Huw V. Smith Scottish Parasite Diagnostic Laboratory, Stobhill Hospital, Glasgow, UK and

R.C. Andrew Thompson WHO Collaborating Centre for the Molecular Epidemiology of Parasitic Infections, School of Veterinary and Biomedical Sciences, Murdoch University, WA, Australia

CABI is a trading name of CAB International CABI Head Office Nosworthy Way Wallingford Oxfordshire OX10 8DE UK Tel: +44 (0)1491 832111 Fax: +44 (0)1491 833508 E-mail: [email protected] Website: www.cabi.org

CABI North American Office 875 Massachusetts Avenue 7th Floor Cambridge, MA 02139 USA Tel: +1 617 395 4056 Fax: +1 617 354 6875 E-mail: [email protected]

© CAB International 2009. All rights reserved. No part of this publication may be reproduced in any form or by any means, electronically, mechanically, by photocopying, recording or otherwise, without the prior permission of the copyright owners. A catalogue record for this book is available from the British Library, London, UK. Library of Congress Cataloging-in-Publication Data Giardia and cryptosporidium : from molecules to disease / edited by Guadalupe Ortega-Pierres . . . [et al.]. p. cm. Includes bibliographical references and index. ISBN 978-1-84593-391-3 (alk. paper) 1. Giardia. 2. Cryptosporidium. 3. Giardiasis–Prevention. I. Ortega-Pierres, Guadalupe. II. C.A.B. International. [DNLM: 1. Giardia. 2. Cryptosporidiosis–prevention & control. 3. Cryptosporidium. 4. Giardiasis–prevention & control. 5. Water Pollution–prevention & control. QX 70 G4346 2008] QL368.D65G53 2008 616.9′3601–dc22 2008025358 ISBN-13: 978 1 84593 391 3 Typeset by AMA Dataset, Preston, UK. Printed and bound in the UK by the MPG Books Group, Bodmin. The paper used for the text pages in this book is FSC certified. The FSC (Forest Stewardship Council) is an international network to promote responsible management of the world's forests.

Contents

Contributors 1

The Impact of Giardia on Science and Society R.C. Andrew Thompson

2

Cryptosporidium in Cattle: From Observing to Understanding Ronald Fayer, Monica Santín and James M. Trout

ix 1

12

TAXONOMY, NOMENCLATURE AND EVOLUTION 3

Names Do Matter Dwight D. Bowman

4

Centenary of the Genus Cryptosporidium: From Morphological to Molecular Species Identification Jan Šlapeta

25

31

MOLECULAR EPIDEMIOLOGY AND TYPING 5

6

7

Molecular Epidemiology of Human Cryptosporidiosis in Developing Countries Lihua Xiao

51

Molecular Epidemiology and Typing of Non-human Isolates of Cryptosporidium Una M. Ryan and Lihua Xiao

65

Insights Into the Molecular Detection of Giardia duodenalis: Implications for Epidemiology Simone M. Cacciò, Marco Lalle, Relja Beck and Edoardo Pozio

81

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ZOONOTIC, HUMAN AND ANIMAL HEALTH ISSUES 8

Wildlife with Giardia: Villain, or Victim and Vector? Susan J. Kutz, R.C. Andrew Thompson and Lydden Polley

9

The Role of Livestock in the Foodborne Transmission of Giardia duodenalis and Cryptosporidium spp. to Humans Brent R. Dixon

10 The Risk of Zoonotic Genotypes of Cryptosporidium spp. in Watersheds Hussni O. Mohammed and Susan E. Wade

94

107

123

CLINICAL AND MOLECULAR EPIDEMIOLOGY 11 Clinical Presentation in Cryptosporidium-infected Patients Laetitia M. Kortbeek 12 Molecular Epidemiology of Cryptosporidium and Giardia Infections Paul R. Hunter

131

138

ADVANCES IN DIAGNOSIS 13 Advances in Diagnosis: is Microscopy Still the Benchmark? 147 Rachel M. Chalmers TREATMENT OF DRINKING WATER 14 Control of Cryptosporidium and Giardia in Surface Water by Disinfection Thomas M. Hargy, Jennifer L. Clancy and Lynn P. Landry

158

15 Towards Methods for Detecting UV-induced Damage in Individual Cryptosporidium parvum and Cryptosporidium hominis Oocysts by Immunofluorescence Microscopy Huw V. Smith, B.H. Al-Adhami, Rosely A.B. Nichols, John R. Kusel and J. O’Grady

179

CONTROL IN WATER 16 Effect of Environmental and Conventional Water Treatment Processes on Waterborne Cryptosporidium Oocysts 198 Brendon King, Alexandra Keegan, Chris Saint and Paul Monis 17 Methods for Genotyping and Subgenotyping Cryptosporidium spp. Oocysts Isolated During Water and Food Monitoring Huw V. Smith, Rosely A.B. Nichols, Lisa Connelly and Christopher B. Sullivan

210

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18 Intervention in Waterborne Disease 227 Gordon Nichols, Iain R. Lake, Rachel M. Chalmers, Graham Bentham, Florence C.D. Harrison, Paul R. Hunter, Sari Kovats, Chris Grundy, Steve Anthony, Hester Lyons, Maureen Agnew and Chris Proctor OTHER WATERBORNE PROTOZOA 19 Occurrence and Control of Naegleria fowleri in Drinking Water Wells Charles P. Gerba, Barbara L. Blair, Payal Sarkar, Kelly R. Bright, R.C. Maclean and Francine Marciano-Cabral 20 Environmental Factors Influencing the Survival of Cyclospora cayetanensis Ynes R. Ortega

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248

BASIC BIOLOGY 21 Recent Advances in the Developmental Biology and Life Cycle of Cryptosporidium 255 Nawal Samih Hijjawi, Annika C. Boxell and R.C. Andrew Thompson 22 Basic Biology of Giardia lamblia: Further Studies on Median Body and Funis 266 Marlene Benchimol 23 Giardia intestinalis: a Microaerophilic Parasite with Mitochondrial Ancestry Gloria León-Avila, José Manuel Hernández and Jorge Tovar

284

METABOLOMICS AND TRANSCRIPTOME 24 Cytoskeleton-based Lipid Transport in a Parasitic Protozoan, Giardia lamblia Cynthia Castillo, Yunuen Hernandez, Sukla Roychowdhury and Siddhartha Das 25 Signalling During Giardia Differentiation Tineke Lauwaet and Frances D. Gillin

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309

GENOMICS 26 Preliminary Analysis of the Cryptosporidium muris Genome 320 Giovanni Widmer, Eric London, Linghui Zhang, Guangtao Ge, Saul Tzipori, Jane M. Carlton and Joana C. da Silva PROTEOMICS 27 Proteomic Analyses in Giardia Daniel Palm and Staffan G. Svärd

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28 Proteomic and Genomic Approaches to Understanding the ‘Power Plant’ of Cryptosporidium Lorenza Putignani, Sanya J. Sanderson, Cristina Russo, Jessica Kissinger, Donato Menichella and Jonathan M. Wastling

344

BIOCHEMISTRY AND PHYSIOLOGY 29 Energy Metabolism and Carbon Flow in Cryptosporidium parvum Guan Zhu

360

30 The Surface Protein Repertoires of Cryptosporidium spp. and Other Apicomplexans Thomas J. Templeton

369

31 Giardan: Structure, Synthesis, Regulation and Inhibition Keriman S ¸ener, Harry van Keulen and Edward L. Jarroll

382

CELL BIOLOGY AND SIGNALLING 32 Protein Kinase C in Giardia duodenalis: a Family Affair M. Luisa Bazán-Tejeda, Raúl Argüello-García, Rosa M. BermúdezCruz, Martha Robles-Flores and Guadalupe Ortega-Pierres 33 Secretory Granule Biogenesis and the Organization of Membrane Compartments via SNARE Proteins in Giardia lamblia Eliana V. Elías, Natalia Gottig, Rodrigo Quiroga and Hugo D. Luján 34 Molecular Mechanisms of Cryptosporidium-induced Host Actin Cytoskeleton Dynamics Steven P. O’Hara, Xian-Ming Chen and Nicholas F. LaRusso

398

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418

PATHOGENESIS AND HOST–PARASITE RELATIONSHIP 35 Pathogenic Mechanisms in Giardiasis and Cryptosporidiosis Andre G. Buret

428

36 Interferon-gamma (IFN-γ) in Immunological Control of Cryptosporidial Infection Naheed Choudhry, Mona Bajaj-Elliott and Vincent McDonald

442

37 Immune Response to Giardia Infection: Lessons from Animal Models Steven M. Singer and Joel Kamda

451

DRUG TREATMENT AND NOVEL DRUG TARGETS 38 Drug Treatment and Novel Drug Targets Against Giardia and Cryptosporidium Jean-François Rossignol

463

Index

483

Contributors

Maureen Agnew, School of Environmental Sciences, University of East Anglia, Norwich, UK. B.H. Al-Adhami, CFIA, Centre for Animal Parasitology, Saskatoon Laboratory, 116 Veterinary Road, Saskatoon, Saskatchewan, Canada S7N 2R3. Steve Anthony, ADAS Consulting, Woodthorne, Wergs Road, Wolverhampton WV6 8TQ, UK. Raúl Argüello-García, Department of Genetics and Molecular Biology, Centro de Investigación y de Estudios Avanzados del IPN, México City, Av. Instituto Politécnico Nacional No. 2508, Col. San Pedro Zacatenco, México, D.F. 07360, Mexico. Mona Bajaj-Elliott, UCL Institute of Child Health, 30 Guilford Street, London WC1N 1EH, UK. M. Luisa Bazán-Tejeda, Department of Genetics and Molecular Biology, Centro de Investigación y de Estudios Avanzados del IPN, México City, Av. Instituto Politécnico Nacional No. 2508, Col. San Pedro Zacatenco, México, D.F. 07360, Mexico. Relja Beck, Department for Parasitology and Parasitic Diseases, University of Zagreb, Heinzelova 55, 10000, Zagreb, Croatia. Marlene Benchimol, Universidade Santa Úrsula, Laboratório de Ultraestrutura Celular, Rua Jornalista Orlando Dantas 59, Botafogo, Rio de Janeiro, R.J., Brazil, CEP 222-31-010. Email: [email protected] Graham Bentham, School of Environmental Sciences, University of East Anglia, Norwich, UK. Rosa M. Bermúdez-Cruz, Department of Genetics and Molecular Biology, Centro de Investigación y de Estudios Avanzados del IPN, México City, Av. Instituto Politécnico Nacional No. 2508, Col. San Pedro Zacatenco, México, D.F. 07360, Mexico. Barbara L. Blair, Department of Soil, Water and Environmental Science, The University of Arizona, Tucson, AZ 85721, USA. ix

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Contributors

Dwight D. Bowman, Department of Microbiology and Immunology, College of Veterinary Medicine, Cornell University, Ithaca, NY 14850, USA. Email: [email protected] Annika C. Boxell, Division of Veterinary and Biomedical Sciences, Murdoch University, Murdoch, WA 6150, Australia. Kelly R. Bright, Department of Soil, Water and Environmental Science, The University of Arizona, Tucson, AZ 85721, USA. Andre G. Buret, Department of Biological Sciences, Inflammation Research Network, University of Calgary, 2500 University Drive N.W., Calgary, Alberta, Canada T2N 1N4. Email: [email protected] Simone M. Cacciò, Department of Infectious, Parasitic and Immunomediated Diseases, Istituto Superiore di Sanità, Viale Regina Elena 299, Rome 00161, Italy. Email: [email protected] Jane M. Carlton, New York University, Department of Medical Parasitology, 550 First Avenue, New York 10016, USA. Cynthia Castillo, Naturopathic Medicine, Bastyr University, 14500 Juanita Drive NE, Kenmore, WA 98028, USA. Rachel M. Chalmers, UK Cryptosporidium Reference Unit (CRU), National Public Health Service for Wales, Microbiology Swansea, Singleton Hospital, Sketty, Swansea SA2 8QA, UK. Email: [email protected] Xian-Ming Chen, Department of Medical Microbiology and Immunology, Creighton University School of Medicine, 2500 California Plaza, Omaha, NE 68178, USA. Naheed Choudhry, Barts and the London School of Medicine and Dentistry, Centre for Gastroenterology, Queen Mary College, University of London, Turner Street, London E1 2AD, UK. Jennifer L. Clancy, Clancy Environmental Consultants, Inc., 20 Mapleville Depot, PO Box 314, Saint Albans, VT 05478, USA. Lisa Connelly, Scottish Parasite Diagnostic Laboratory (SPDL), Stobhill Hospital, 133 Balornock Road, Glasgow G21 3UW, UK. Siddhartha Das, Infectious Diseases and Immunology Unit, The Border Biomedical Research Center, Department of Biological Sciences, University of Texas at El Paso, 500 W. University Avenue, El Paso, TX 79968, USA. Email: [email protected] Brent R. Dixon, Microbiology Research Division, Banting Research Centre, 251 Sir Frederick Banting Driveway, PL 2204A2, Ottawa, Ontario, Canada K1A 0K9. Email: [email protected] Eliana V. Elías, Mercedes & Martin Ferreyra Institute for Medical Research, INIMEC-CONICET and School of Medicine, Catholic University of Cordoba, Parque Velez Sarsfield, CP 5000, Cordoba, Argentina. Ronald Fayer, Environmental Microbial Safety Laboratory, Animal and Natural Resources Institute, Agricultural Research Service, United States Department of Agriculture, Building 173, BARC-East, 10300 Baltimore Avenue, Beltsville, MD 20705, USA. Email: [email protected] Guangtao Ge, Department of Computer Sciences, School of Engineering, Tufts University, 200 Westboro Road, North Grafton, MA 01536, USA.

Contributors

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Charles P. Gerba, Department of Soil, Water and Environmental Science, The University of Arizona, Tucson, AZ 85721, USA. Email: [email protected]. edu Frances D. Gillin, Department of Pathology, Division of Infectious Diseases, University of California at San Diego, San Diego, CA, USA. Email: [email protected] Natalia Gottig, Mercedes & Martin Ferreyra Institute for Medical Research, INIMEC-CONICET and School of Medicine, Catholic University of Cordoba, Parque Velez Sarsfield, CP 5000, Cordoba, Argentina. Chris Grundy, Public and Environmental Health Research Unit, London School of Hygiene and Tropical Medicine, Keppel Street, London, WC1E 7HT, UK. Thomas M. Hargy, Clancy Environmental Consultants, Inc., 20 Mapleville Depot, PO Box 314, Saint Albans, VT 05478, USA. Florence C.D. Harrison, School of Environmental Sciences, University of East Anglia, Norwich, UK. José Manuel Hernández, Department of Cell Biology, Centro de Investigación y de Estudios Avanzados del IPN, Av. IPN 2508, San Pedro, Zacatenco, CP 07300, México D.F., Mexico. Yunuen Hernandez, Laboratory of Parasitic Diseases, National Institutes of Allergy and Infectious Diseases, Bldg 4/Rm16, 4 Center Drive, Bethesda, MD 20892, USA. Nawal Samih Hijjawi, Department of Medical Laboratory Sciences, Faculty of Allied Health Sciences, The Hashemite University, PO Box 150459, Zarqa 13115, Jordan. Paul R. Hunter, School of Medicine, Health Policy and Practice, University of East Anglia, Earlham Road, Norwich NR4 7TJ, UK. Email: Paul.Hunter@ uea.ac.uk Edward L. Jarroll, Department of Biology, Northeastern University, Boston, MA 02115, USA. Email: [email protected] Joel Kamda, Department of Biology and Center for Infectious Diseases, Georgetown University, Washington, DC 20057, USA. Alexandra Keegan, The Co-operative Research Centre for Water Quality and Treatment, Australian Water Quality Centre, SA Water Corporation, Salisbury, South Australia 5108, Australia. Brendon King, The Co-operative Research Centre for Water Quality and Treatment, Australian Water Quality Centre, SA Water Corporation, Salisbury, South Australia 5108, Australia. Jessica Kissinger, Center for Tropical and Emerging Global Diseases and Department of Genetics, University of Georgia, 500 D.W. Brooks Drive, Athens, GA 30602, USA. Laetitia M. Kortbeek, Centre for Infectious Disease Control Netherlands, Laboratory for Infectious Diseases and Perinatal Screening (LIS), National Institute of Public Health and the Environment, PO Box 1, 3720 BA Bilthoven, the Netherlands. Email: [email protected] Sari Kovats, Public and Environmental Health Research Unit, London School of Hygiene and Tropical Medicine, Keppel Street, London WC1E 7HT, UK.

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Contributors

John R. Kusel, Division of Infection and Immunity, Institute of Biomedical and Life Sciences, University of Glasgow, Glasgow G12 8QQ, UK. Susan J. Kutz, Faculty of Veterinary Medicine, University of Calgary, 3330 Hospital Drive NW, Calgary, Alberta, Canada T2N 4N1. Email: [email protected] Iain R. Lake, School of Environmental Sciences, University of East Anglia, Norwich, UK. Marco Lalle, Department of Infectious, Parasitic and Immunomediated Diseases, Istituto Superiore di Sanità, Viale Regina Elena 299, Rome 00161, Italy. Lynn P. Landry, Utility Analysis and Environmental Management, Policy and Planning Department, Metro Vancouver, Greater Vancouver Regional District, 4330 Kingsway, Burnaby, BC, Canada V5H 4G8. Nicholas F. LaRusso, Miles and Shirley Fiterman Center for Digestive Diseases, Division of Gastroenterology and Hepatology, Mayo Clinic College of Medicine, 200 First Street SW, Rochester, MN 55905, USA. Email: larusso. [email protected] Tineke Lauwaet, Department of Pathology, Division of Infectious Diseases, University of California at San Diego, San Diego, CA, USA. Gloria León-Avila, Department of Parasitology, Escuela Nacional de Ciencias Biológicas, IPN, Carpio y Plan de Ayala, Casco de Santo Tomás, CP 11340, México, D.F., Mexico. Email: [email protected] Eric London, Tufts Cummings School of Veterinary Medicine, Division of Infectious Diseases, Tufts University, 200 Westboro Road, North Grafton, MA 01536, USA. Hugo D. Luján, Mercedes & Martin Ferreyra Institute for Medical Research, INIMEC-CONICET and School of Medicine, Catholic University of Cordoba, Parque Velez Sarsfield, CP 5000, Cordoba, Argentina. Email:hlujan@ immf.uncor.edu Hester Lyons, ADAS Consulting, Woodthorne, Wergs Road, Wolverhampton WV6 8TQ, UK. R.C. Maclean, Department of Microbiology and Immunology, Medical College of Virginia Campus, Virginia Commonwealth University, Richmond, VA 23298, USA. Francine Marciano-Cabral, Department of Microbiology and Immunology, Medical College of Virginia Campus, Virginia Commonwealth University, Richmond, VA 23298, USA. Vincent McDonald, Barts and the London School of Medicine and Dentistry, Centre for Gastroenterology, Queen Mary College, University of London, Turner Street, London E1 2AD, UK. Email: [email protected] Donato Menichella, Unit of Microbiology and Virology, Bambino Gesù Hospital, Scientific Institute, Piazza Sant’Onofrio 4, 00146, Rome, Italy. Email: [email protected] Hussni O. Mohammed, Department of Population Medicine and Diagnostic Sciences, College of Veterinary Medicine, Cornell University, Ithaca, NY 14853, USA. Email: [email protected] Paul Monis, The Co-operative Research Centre for Water Quality and Treatment, Australian Water Quality Centre, SA Water Corporation, Salisbury, South Australia 5108, Australia. Email:[email protected]

Contributors

xiii

Gordon Nichols, Environmental and Enteric Diseases Department, Communicable Disease Surveillance Centre, Health Protection Agency (HPA) Centre for Infections, 61 Colindale Avenue, London NW9 5EQ, UK. Email: [email protected] Rosely A.B. Nichols, Scottish Parasite Diagnostic Laboratory (SPDL), Stobhill Hospital, 133 Balornock Road, Glasgow G21 3UW, UK. J. O’Grady, Strathclyde Institute of Pharmacy and Biomedical Sciences, University of Strathclyde, 27 Taylor Street, Glasgow G4 0NR, UK. Steven P. O’Hara, Miles and Shirley Fiterman Center for Digestive Diseases, Division of Gastroenterology and Hepatology, Mayo Clinic College of Medicine, 200 First Street SW, Rochester, MN 55905, USA. Ynes R. Ortega, Center for Food Safety, University of Georgia, 1109 Experiment Street, Griffin, GA 30223, USA. Email: [email protected] Guadalupe Ortega-Pierres, Department of Genetics and Molecular Biology, Centro de Investigación y de Estudios Avanzados del IPN, México City, Av. Instituto Politécnico Nacional No. 2508, Col. San Pedro Zacatenco, México, D.F. 07360, Mexico. Email: [email protected] Daniel Palm, Centre for Microbial Preparedness, Swedish Institute for Infectious Disease Control, SE-171 82 Solna, Sweden. Lydden Polley, Western College of Veterinary Medicine, University of Saskatchewan, 52 Campus Drive, Saskatoon, SK, Canada S7N 5B4. Email:lydden. [email protected] Edoardo Pozio, Department of Infectious, Parasitic and Immunomediated Diseases, Istituto Superiore di Sanità, Viale Regina Elena 299, Rome 00161, Italy. Chris Proctor, ADAS Consulting, Woodthorne, Wergs Road, Wolverhampton WV6 8TQ, UK. Lorenza Putignani, Unit of Microbiology and Virology, Bambino Gesù Hospital, Scientific Institute, Piazza Sant’Onofrio 4, 00146, Rome, Italy. Email: [email protected] Rodrigo Quiroga, Mercedes & Martin Ferreyra Institute for Medical Research, INIMEC-CONICET and School of Medicine, Catholic University of Cordoba, Parque Velez Sarsfield, CP 5000, Cordoba, Argentina. Martha Robles-Flores, Departamento de Bioquímica, Facultad de Medicina, Universidad Nacional Autónoma de México, Apdo. Postal 70-159, México City, D.F. 04510, Mexico. Jean-François Rossignol, Division of Gastroenterology and Hepatology, Department of Medicine, Stanford University School of Medicine, 300 Pasteur Drive, Stanford, CA 94305, USA. Email: [email protected] Sukla Roychowdhury, Infectious Diseases and Immunology Unit, The Border Biomedical Research Center, Department of Biological Sciences, University of Texas at El Paso, 500 W. University Avenue, El Paso, TX 79968, USA. Cristina Russo, Unit of Microbiology and Virology, Bambino Gesù Hospital, Scientific Institute, Piazza Sant’Onofrio 4, 00146, Rome, Italy. Email:russocri@ opbg.net Una M. Ryan, School of Veterinary and Biomedical Sciences, Murdoch University, South Street, Murdoch, WA 6150, Australia. Email: Una.Ryan@ murdoch.edu.au

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Contributors

Chris Saint, The Co-operative Research Centre for Water Quality and Treatment, Australian Water Quality Centre, SA Water Corporation, Salisbury, South Australia 5108, Australia. Sanya J. Sanderson, Department of Pre-Clinical Veterinary Science and Veterinary Pathology, Faculty of Veterinary Science, University of Liverpool, Crown Street, Liverpool L69 7ZJ, UK. Email: [email protected] Monica Santín, Environmental Microbial Safety Laboratory, Animal and Natural Resources Institute, Agricultural Research Service, United States Department of Agriculture, Building 173, BARC-East, 10300 Baltimore Avenue, Beltsville, MD 20705, USA. Payal Sarkar, Department of Soil, Water and Environmental Science, The University of Arizona, Tucson, AZ 85721, USA. Keriman S¸ener, Department of Biology, Northeastern University, Boston, MA 02115, USA. Email:[email protected] Joana C. da Silva, Institute for Genome Sciences, University of Maryland School of Medicine, 20 Penn Street, Baltimore, MD 21201, USA. Steven M. Singer, Department of Biology and Center for Infectious Diseases, Georgetown University, Washington, DC 20057, USA. Email: sms3@ georgetown.edu Jan Šlapeta, Faculty of Veterinary Science, McMaster Building B14, University of Sydney, NSW 2006, Australia. Email: [email protected] Huw V. Smith, Scottish Parasite Diagnostic Laboratory (SPDL), Stobhill Hospital, 133 Balornock Road, Glasgow G21 3UW, UK. Email: huw.smith@ northglasgow.scot.nhs.uk Christopher B. Sullivan, Scottish Parasite Diagnostic Laboratory (SPDL), Stobhill Hospital, 133 Balornock Road, Glasgow G21 3UW, UK. Staffan G. Svärd, Department of Cell and Molecular Biology, BMC, Uppsala University, Box 596, SE-751 24 Uppsala, Sweden. Email: staffan.svard@ icm.uu.se Thomas J. Templeton, Department of Microbiology and Immunology, Weill Medical College of Cornell University, 1300 York Avenue, Box 62, NY 10021 USA. Email: [email protected] R.C. Andrew Thompson, WHO Collaborating Centre for the Molecular Epidemiology of Parasitic Infections, Division of Veterinary and Biomedical Sciences, Murdoch University, Murdoch, WA 6150, Australia. Email: a.thompson@ murdoch.edu.au Jorge Tovar, School of Biological Sciences, Royal Holloway University of London, Egham TW20 0EX, UK. James M. Trout, Environmental Microbial Safety Laboratory, Animal and Natural Resources Institute, Agricultural Research Service, United States Department of Agriculture, Building 173, BARC-East, 10300 Baltimore Avenue, Beltsville, MD 20705, USA. Saul Tzipori, Tufts Cummings School of Veterinary Medicine, Division of Infectious Diseases, Tufts University, 200 Westboro Road, North Grafton, MA 01536, USA. Harry van Keulen, Department of Biological, Geological and Environmental Sciences, Cleveland State University, Cleveland, OH 44115, USA.

Contributors

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Susan E. Wade, Department of Population Medicine and Diagnostic Sciences, College of Veterinary Medicine, Cornell University, Ithaca, NY 14853, USA. Jonathan M. Wastling, Department of Pre-Clinical Veterinary Science and Veterinary Pathology, Faculty of Veterinary Science, University of Liverpool, Crown Street, Liverpool L69 7ZJ, UK. Email: [email protected] Giovanni Widmer, Tufts Cummings School of Veterinary Medicine, Division of Infectious Diseases, Tufts University, 200 Westboro Road, North Grafton, MA 01536, USA. Email: [email protected] Lihua Xiao, Division of Parasitic Diseases, National Center for Infectious Diseases, Centers for Disease Control and Prevention, Building 22, Mail Stop F-12, 4770 Buford Highway, Atlanta, GA 30341, USA. Email: [email protected] Linghui Zhang, Tufts Cummings School of Veterinary Medicine, Division of Infectious Diseases, Tufts University, 200 Westboro Road, North Grafton, MA 01536, USA. Guan Zhu, Department of Veterinary Pathobiology, College of Veterinary Medicine and Biomedical Sciences, Texas A&M University, 4467 TAMU, College Station, TX 77843, USA. Email: [email protected]

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1

The Impact of Giardia on Science and Society R.C.A. THOMPSON WHO Collaborating Centre for the Molecular Epidemiology of Parasitic Infections, Murdoch University, WA, Australia

Abstract Although Giardia has a long history, it is only recently that the clinical impact of Giardia in children has been recognized. Similarly, the emergence of Giardia as a frequent parasite of companion animals, livestock and wildlife raises questions about the clinical and zoonotic significance of such infections. Transmission patterns have already been described but the frequency of interaction between cycles of transmission is only just beginning to be addressed in molecular epidemiological studies. The application of molecular tools has provided information which sets the scene for revising the taxonomy of Giardia. With the completion of the Giardia genome sequencing project, it is hoped that it will soon be possible to compare genome and phenome and provide information of practical value for the control of Giardia, as well as increasing our understanding of the evolution and phylogenetic relationships of Giardia.

Introduction The protozoa that collectively comprise the genus Giardia have intrigued biologists and clinicians for over 300 years, ever since Antony van Leeuwenhoek first discovered the organism (Meyer, 1994). This enigmatic protozoan possesses a number of unusual characteristics including the presence of two similar, transcriptionally active diploid nuclei, the absence of mitochondria and peroxisomes, and a unique attachment organelle – the ventral sucking disc (Thompson and Monis, 2004; Morrison et al., 2007). Phylogenetic relationships are controversial, with one school of thought suggesting that Giardia is a basal eukaryote and the other that Giardia comprises one of many divergent eukaryotic lineages that adapted to a microaerophilic lifestyle rather than diverging before the endosymbiosis of the mitochondrial ancestor (Thompson and Monis, 2004; Morrison et al., 2007). Despite its long history, our understanding of the pathogenesis of Giardia infections and its relationship with its host is limited, and we do not know why clinical disease occurs in some individuals but may not be apparent in others © CAB International 2009. Giardia and Cryptosporidium: From Molecules to Disease (eds G. Ortega-Pierres et al.)

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(Savioli et al., 2006). There are no known virulence factors or toxins, and variable expression of surface proteins may allow evasion of host immune responses and adaptation to different environments (Morrison et al., 2007). For many years after van Leeuwenhoek first described Giardia, there was controversy as to whether it was a parasite that caused disease or a commensal. It was not until the 1920s that influential proponents such as Dobell and Miller gave support to the link between the presence of Giardia and disease and, more importantly, a link with malabsorption syndromes (Cox, 1998). Thirty years later, Rendtorff established infections with Giardia in humans following the oral inoculation of cysts (Rendtorff, 1954). Today, there is no doubt about the clinical significance of Giardia infections in humans, and particularly the impact of giardiasis in children and its association with failure to thrive and wasting syndromes (Thompson, 2008).

Taxonomy The recent application of molecular, PCR-based tools has enabled the genetic relationships of a range of morphologically identical ‘strains’ of Giardia to be determined (Thompson and Monis, 2004; Cacciò et al., 2005; Traub et al., 2005; Smith et al., 2006). As a consequence, a large number of species and genotypes of Giardia are now recognized that differ principally in their host range. The current taxonomy of Giardia is summarized in Table 1.1 and has been extensively reviewed (Thompson and Monis, 2004; Cacciò et al., 2005; Thompson et al., 2007). The nomenclature most widely accepted at the present time for the genotypes that have been characterized is ‘assemblage’, although a revised taxonomy has been proposed (Thompson and Monis, 2004; Cacciò et al., 2005). Some species and genotypes/assemblages appear to be restricted to particular species or types of hosts (e.g. Giardia assemblages C/D (G. canis) and E (G. bovis) in dogs and livestock, respectively; see Table 1.1) whereas others have broad host ranges, including humans, (e.g. G. duodenalis assemblages A and B; see Table 1.1) and are therefore of zoonotic significance. Giardia duodenalis (syn. G. intestinalis, G. lamblia) is the only species found in humans. Table 1.1 summarizes the proposed revised taxonomy for assemblages of Giardia. Such a formal nomenclature will avoid confusion and enhance communication.

Clinical Impact Humans In developed countries, infections with Giardia are most common in children, especially in daycare centres and among travellers, and a rising incidence in such settings has led to the designation of giardiasis as a re-emerging infectious disease in the developed world (Thompson, 2000, 2004; Eckmann, 2003; Thompson and Monis, 2004). The World Health Organization (WHO) has given consideration to intestinal protozoa for many years, but because of their very different disease dynamics they did not initially form part of the ‘Neglected Diseases Initiative’.

The Impact of Giardia on Science and Society Table 1.1.

3

Established and proposeda species/assemblages of Giardia.

Species/assemblage

Host

G. duodenalis/assemblage A

Humans and other primates, dogs, cats, livestock, rodents and other wild mammals Humans and other primates, dogs

G. duodenalis/assemblage B (G. enterica)a G. agilis G. muris G. psittaci G. ardeae G. duodenalis/assemblage C/D (G. canis)a G. duodenalis/assemblage F (G. cati)a G. duodenalis/assemblage E (G. bovis)a G. duodenalis/assemblage G (G. simondi )a aSee

Amphibians Rodents Birds Birds Dogs Cats Cattle and other hoofed livestock Rats

Thompson and Monis (2004), Cacciò et al. (2005).

However, since they all have a common link with poverty, the current view is to take a comprehensive approach to all these diseases. In September 2004, Giardia was included in WHO’s Neglected Diseases Initiative (Savioli et al., 2006). In developing countries, particularly in Asia, Africa and Latin America, about 200 million people have symptomatic giardiasis with some 500,000 new cases being reported each year (WHO, 1996). Children living in communities are most commonly infected in developing countries, particularly among disadvantaged groups living in isolated communities, such as Australian Aborigines (Thompson, 2000; Hesham et al., 2005; Savioli et al., 2006). These children are most at risk from the chronic consequences of Giardia infection. Although animals may serve as reservoirs of Giardia infection that under certain circumstances may spill over to humans; from a clinical viewpoint, direct human-tohuman transmission is of most significance, particularly in situations where the frequency of transmission is high. Human-to-human transmission of Giardia can occur indirectly through the accidental ingestion of cysts in contaminated water or food, or directly in environments where hygiene levels may be compromised, such as daycare centres or disadvantaged community settings, where the frequency of transmission is high and/or conditions are conducive to direct person-to-person transfer (Thompson, 2000; Hesham et al., 2005). Under such circumstances, children may be at constant risk of infection, even though chemotherapeutic interventions may be instituted (Thompson et al., 2001; Savioli et al., 2006). If children are constantly exposed they will be re-infected rapidly, since anti-giardial agents have no residual activity. The fact that children in such endemic settings do not appear to develop resistance to Giardia infection may be due to suboptimal immunological competence and/or infection with different ‘strains’/subgenotypes of Giardia (Hopkins et al.,

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1999). It might be expected that competitive interactions would result in the predominance of particular genotypes of Giardia and the exclusion of others (Thompson et al., 2001), but this does not appear to be the case, perhaps due to the suboptimal nutritional and immunological environments to which the parasites are exposed in such communities. Symptomatic infection in humans may not be evident in a significant proportion of infected individuals (Troeger et al., 2007), and represents only a fraction (20–80%) of all stool-positive Giardia infections (Nash et al., 1987; Flanagan, 1992; Rodriguez-Hernandez et al., 1996). Symptoms are highly variable but include persistent, usually short-term, diarrhoea, epigastric pain, nausea, vomiting and weight loss (Thompson et al., 1993; Eckmann, 2003). Symptoms typically occur 6–15 days after infection and last for 2–4 days. As such, infection is assumed to be self-limiting in more than 85% of cases (indicating that effective host defences exist), although chronic cases occur occasionally in the absence of apparent immunodeficiencies (Nash et al., 1987; Flanagan, 1992). The risk factors for clinical giardiasis, particularly in humans, have yet to be resolved, but clearly involve host and environmental factors as well as the ‘strain’/ genotype/assemblage of the parasite (Buret, 2007). However, a distinction needs to be made between the effects of a single infection, which may give rise to the ‘classical’ short-term episode of diarrhoea, and the long-term effects of persistent Giardia infection, particularly in children, in environments where the frequency of transmission is high. Here the picture is very different. In endemic foci where the frequency of transmission is high and often enhanced by poor hygiene and environmental contamination, children are at particular risk from the more serious and long-term consequences of Giardia infection that are associated with malnutrition, micronutrient deficiency and failure to thrive, iron deficiency anaemia and poor cognitive function (Hesham et al., 2004; Savioli et al., 2006; Gbakima et al., 2007; Gonen et al., 2007). Clearly, the impact of Giardia in such circumstances will be exacerbated by poor/suboptimal nutrition and concurrent infections with other enteric parasites such as Hymenolepis nana, Entamoeba coli and Blastocystis. Longitudinal studies on the impact of enteric parasites on childhood growth and mental development in such endemic areas is urgently required (Savioli et al., 2006). The children who are infected in such environments, particularly in developing countries and among disadvantaged groups, represent the most important group in terms of the clinical impact of Giardia (Thompson, 2000; Hesham et al., 2005; Savioli et al., 2006). In such circumstances, it is debatable whether the regular use of drugs is of any benefit. This is in contrast to the situation with gastrointestinal helminths such as hookworm, where regular mass chemotherapy has been shown to have great benefit in control (Reynoldson et al., 1998; Thompson et al., 2001).

Domestic animals Livestock, wildlife, and companion animals are frequently infected with Giardia and are susceptible to host-adapted and zoonotic species of Giardia. Although infections with Giardia are common in dogs and cats, most infected animals

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remain asymptomatic. If clinical disease occurs, it is usually associated with young animals and those in kennel or cattery situations (Robertson et al., 2000), where the effects of overcrowding, weaning and nutritional deficiency, may cause stress and exacerbate the effects of an infection (Thompson, 2004). The most consistent clinical sign of giardiasis in dogs and cats is small-bowel diarrhoea, which may be acute or chronic, and self-limiting, intermittent or continuous in nature. Giardia infection in ruminants is often asymptomatic but may also be associated with the occurrence of diarrhoea and ill-thrift in calves (O’Handley et al., 1999; Geurden et al., 2006). The importance of giardiasis as a cause of diarrhoea in ruminants is unclear, especially given that diarrhoea in ruminants is often multifactorial, with more than one pathogen detected (O’Handley and Olson, 2006). Nevertheless, the significance of Giardia infection in ruminants warrants further investigation, particularly with regard to production loss. In cattle, Giardia is commonly found alone or in combination with other pathogens as a cause of calf diarrhoea, which can have economic significance (O’Handley et al., 1999; Olson et al., 2004). In two studies in sheep, Giardia reduced the rate of weight gain, impaired feed efficiency and decreased carcass weight (Olson et al., 1995; Aloisio et al., 2006).

Wildlife It is often a common ‘knee-jerk’ reaction when parasites with zoonotic potential are found in wildlife that they represent a threat to public health as a reservoir and potential source of infection for humans (Thompson, 2004). Indeed, this was the case when WHO initially listed the common enteric protozoan parasite Giardia as a zoonosis over 25 years ago as a result of epidemiological observations suggesting that giardiasis in campers in Canada was caused by drinking stream water contaminated with Giardia from beavers (Thompson, 2004). No one thought to ask the question of where the beavers got their Giardia infections from until only beavers living downstream from a sewage works were found to be infected. With the subsequent application of molecular tools, it has been confirmed that beavers are susceptible to zoonotic strains of Giardia (see Thompson, 2004). The question now is: are they victim or villain with respect to human giardiasis? A similar situation has been reported in non-human primates for which there is a growing literature of the invasion of human pathogens into wild populations (Sleeman et al., 2000; Graczyk et al., 2001, 2002). For example, it was suggested that the discovery of Giardia and the cohabiting enteric protozoan Cryptosporidium in mountain gorillas in the Bwindi Impenetrable National Park, Uganda, was thought to indicate enhanced contact with humans and/or domestic livestock. This was confirmed when rangers and their cattle were found to be infected with Giardia and that the genotype was the same as that recovered from the gorillas (Graczyk et al., 2002). Muskoxen (Ovibos moschatus) are indigenous to the arctic tundra of Canada and Greenland and have been translocated to areas in Alaska, the USA, Russia, Norway and Sweden. These animals are well adapted to their northern environment, and tend to have a relatively simple parasite fauna. Recent surveys on the

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biodiversity and impacts of parasites in Arctic ungulates described Giardia duodenalis, assemblage A, the zoonotic genotype, in muskoxen (Kutz et al., 2008). This unexpected finding (a novel strain, or the livestock strain, was predicted) raises many interesting questions regarding the origin and epidemiology of this parasite in humans and wildlife in this Arctic ecosystem. In particular, is this a pathogen initially introduced to muskoxen by humans? Is Giardia now maintained as a sylvatic cycle in muskoxen (or other wildlife species in the locality) independent of humans? Does the Giardia from muskoxen spill back into humans? The impact of Giardia on the health and production (body condition, fecundity and pelage) of free-ranging ungulates, including muskoxen, remains unknown. In Australia, marsupials are commonly infected with Giardia (Kettlewell et al., 1998) but until recently it was not known to what species or strain(s) of Giardia they were susceptible. Studies on the quenda (Isoodon obesulus), a common widespread species of bandicoot in southern Australia, demonstrated that they were infected with a novel, genetically distinct, form of Giardia, so different from what has been described from humans and other animals that it probably represents a distinct species (Adams et al., 2004). The Giardia isolates genotyped from quenda in their natural habitats have all proved to be the novel strain. However, when quenda were trapped and examined on a farm, they were found to be infected with ‘domestic’ strains of Giardia normally found in livestock and humans (from assemblages A and E; see Table 1.1). Presumably, this reflects the susceptibility of the quenda to other strains of Giardia, as with the case of beavers in North America. This case study raises questions regarding the pathogenicity of non-host-adapted strains of Giardia in naïve wildlife hosts. Additionally, it also raises the question of competition between cohabiting ‘strains’ of Giardia (Thompson and Monis, 2004) and whether, in this case, and perhaps in other species of wildlife, zoonotic strains of Giardia can out-compete the host-specific wildlife strains.

Molecular Epidemiology Humans may be infected with Giardia genotypes belonging to assemblage A or assemblage B (Thompson and Monis, 2004; Cacciò et al., 2005). There is considerable evidence of phenotypic differences between these two assemblages in characters such as metabolism and growth rate (Thompson and Monis, 2004). It has therefore been proposed that there may be differences in the nature of infection with these two assemblages in humans which may be reflected in duration of infection, drug sensitivity and virulence (Thompson and Monis, 2004). There is growing evidence to support these suggestions but there is a need for more focused molecular epidemiological studies. For example, in tea-growing communities in Assam, India, the proportion of assemblage B and A infections in 18 infected people was 61% and 39%, respectively (Traub et al., 2004). Another study in the UK that examined 35 human clinical samples found that 64% were assemblage B, 27% were assemblage A genetic subgroup II, and the remainder were a mixture of assemblage B and assemblage A genetic group II (Amar et al., 2002).

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There is also increasing evidence supporting differences in virulence between genetic groups of G. duodenalis. Several studies have examined the relationship between clinical symptoms and the genetic assemblage of G. duodenalis infecting human patients. Three of these, undertaken in Australia, Bangladesh and Spain, all found a statistically significant correlation between the presence of symptoms, as defined by diarrhoea, and infection with Giardia belonging to assemblage A (Read et al., 2002; Haque et al., 2005; Sahagún et al., 2007). In contrast, it was found, in all studies, that infections with assemblage B were usually asymptomatic. A study in the Netherlands of individuals presenting general practitioners with diarrhoeal complaints found that those infected with assemblage A isolates had intermittent diarrhoea, whereas those infected with assemblage B had acute or persistent diarrhoea (Homan and Mank, 2001). However, this study only sampled patients presenting with diarrhoeal complaints. There is a need for additional large-scale molecular epidemiological surveys of Giardia infections in humans. With the limited data currently available it is not possible to determine the geographical distribution and prevalence of human-infective genotypes. With such data it may be possible to determine the significance of any ‘strain’-related differences in virulence. Although studies on the occurrence of the different genotypes of Giardia serve to emphasize the potential public health risk from domestic dogs and cats, data on the frequency of zoonotic Giardia transmission are lacking (Thompson, 2004; Leonhard et al., 2007). Such information can be obtained from molecular epidemiological studies that genotype isolates of the parasites from susceptible hosts in localized endemic foci of transmission or as a result of longitudinal surveillance and genotyping of positive cases. In the former, recent research in localized endemic foci of transmission has provided evidence in support of the role of dogs in cycles of zoonotic Giardia transmission involving humans and domestic dogs from communities in tea-growing areas of Assam, India, and in temple communities in Bangkok, Thailand (Traub et al., 2004; Inpankaew et al., 2007). In both these studies, some dogs and their owners sharing the same living area were shown to harbour isolates of G. duodenalia from the same assemblage. Other studies have shown that zoonotic genotypes of Giardia may occur frequently in individual pet dogs living in urban areas (for a review, see Leonhard et al., 2007).

Future Perspectives Sex Population genetic studies of Giardia in communities where the frequency of transmission is very high, have found evidence of occasional bouts of genetic exchange in Giardia (Meloni et al., 1995). These authors demonstrated multiple banding patterns in a number of isolates of Giardia by allozyme electrophoresis which, if a true reflection of the underlying genotypes of the isolates, would seem to indicate that G. duodenalis is functionally diploid, and that recombination or sexual reproduction must have occurred at some stage to produce the apparent

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heterozygotes (Tait, 1983; Meloni et al., 1995). These observations have been supported by subsequent population genetic studies (Cooper et al., 2007) and recent genomic studies (Ramesh et al., 2005; Morrison et al., 2007; Teodorovic et al., 2007). The evolutionary advantage this gives to Giardia is the capacity to respond to adversity, for example selection pressures imposed by regular exposure to anti-giardial drugs or competition with cohabiting ‘strains’ in circumstances where the likelihood of mixed infections is common (Hopkins et al., 1999).

Correlating phenotype and genotype Genetic studies and sequencing of the Giardia genome have laid an important foundation for understanding this parasite. As a consequence of delineating genotypic groupings, a clearer picture of transmission patterns and host specificity has been obtained. However, phenotypic differences in virulence, drug sensitivity and infectivity have been reported in isolates of G. duodenalis and there is a need to correlate these observations with genetic differences (Thompson and Monis, 2004; Cacciò et al., 2005). The correlation of phenotype and genotype will provide important information about the host–parasite relationship in Giardia infections. The complexity of any biological system, including that of Giardia, lies at the protein level and genomics alone cannot be used to understand these complexities. Using a proteomic approach to examine differences between the human infective genotypes of Giardia, we have found differences in a number of proteins between the two human infective genetic groups – assemblages A and B. These proteins of difference appear to be associated with virulence and pathogenicity but at what level of functionality remains unclear at this stage (Steuart et al., in press). We have also identified proteins, some of which are novel and Giardia-specific, that appear to be key to its survival and transmission, including growth factors and developmental triggers. Giardia is also of evolutionary and biological significance in terms of understanding the origin of higher animals from bacteria as well as fundamental questions about the parasitic way of life. Thus some of the proteins we have identified may contribute to a better understanding of Giardia’s pivotal position in our understanding of eukaryote biology and evolution.

References Adams, P.J., Monis, P.T., Elliot, A.D. and Thompson, R.C. (2004) Cyst morphology and sequence analysis of the small subunit rDNA and ef1 alpha identifies a novel Giardia genotype in a quenda (Isoodon obesulus) from Western Australia. Infection, Genetics and Evolution 4, 365–370. Aloisio, F., Filippini, G., Antenucci, P., Lepri, E., Pezzotti, G., Cacciò, S.M. and Pozio, E. (2006) Severe weight loss in lambs infected with Giardia duodenalis assemblage B. Veterinary Parasitology 142, 154–158. Amar, C.F.L., Dear, P.H., Pedraza-Díaz, S., Looker, N., Linnane, E. and McLauchlin, J. (2002) Sensitive PCR-restriction fragment length polymorphism assay for detection

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and genotyping of Giardia duodenalis in human feces. Journal of Clinical Microbiology 40, 446–452. Buret, A.G. (2007) Mechanisms of epithelial dysfunction in giardiasis. Gut 56, 316–317. Cacciò, S.M., Thompson, R.C.A., McLauchlin, J. and Smith, H.V. (2005) Unravelling Cryptosporidium and Giardia epidemiology. Trends in Parasitology 21, 430–437. Cooper, M.A., Adam, R.D., Worobey, M. and Sterling, C.R. (2007) Population genetics provides evidence for recombination in Giardia. Current Biology 17, 1984–1988. Cox, F. (1998) History of human parasitology. In: Cox, F.E.G., Kreier, J.P. and Wakelin, D. (eds) Microbiology and Microbial Infections. Vol. 5: Parasitology. Arnold, London, pp. 3–18. Eckmann, L. (2003) Mucosal defences against Giardia. Parasite Immunology 25, 259–270. Flanagan, P.A. (1992) Giardia – diagnosis, clinical course and epidemiology: a review. Epidemiology and Infection 109, 1–22. Gbakima, A.A., Konteh, R., Kallon, M., Mansaray, H., Sahr, F., Bah, Z.J., Spencer, A. and Luckay, A. (2007) Intestinal protozoa and intestinal helminthic infections in displacement camps in Sierra Leone. African Journal of Medical Science 36, 1–9. Gonen, C., Yilmaz, N., Yalcin, M., Simsek, I. and Gonen, O. (2007) Diagnostic yield of routine duodenal biopsies in iron deficiency anaemia: a study from Western Australia. European Journal of Gastroenterology and Hepatology 19, 37–41. Geurden, T., Claerebout, E., Dursin, L., Deflandre, A., Bernay, F., Kaltsatos, V. and Vercruysse, J. (2006) The efficacy of an oral treatment with paromomycin against an experimental infection with Giardia in calves. Veterinary Parasitology 135, 241–247. Graczyk, T.K., DaSilva, A.J., Cranfield, M.R., Nizeyi, J.B., Kalema, G.R.N.N. 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. Parasitology Research 87, 368–370. Graczyk, T.K., Bosco-Nizeyi, J., Ssebide, B., Thompson, R.C., Read, C. and Cranfield, M.R. (2002) Anthropozoonotic Giardia duodenalis genotype (assemblage) A infections in habitats of free-ranging human-habituated gorillas, Uganda. Journal of Parasitology 88, 905–909. Haque, R., Roy, S., Kabir, M., Stroup, S.E., Mondal, D. and Houpt, E.R. (2005) Giardia assemblage A infection and diarrhoea in Bangladesh. Journal of Infectious Diseases 192, 2171–2173. Hesham, M.S., Edariah, A.B. and Norhavat, M. (2004) Intestinal parasitic infections and micronutrient deficiency: a review. Medical Journal of Malaysia 59, 284–293. Hesham, M.S., Azlin, M., Nor Aini, U.N., Shaik, A., Sa’iah, A., Fatmah, M.S., Ismail, M.G., Firdaus, M.S.A., Aisah, M.Y., Rozlida, A.R. and Norhayati, M. (2005) Giardiasis as a predictor of childhood malnutrition in Orang Asli children in Malaysia. Transactions of the Royal Society for Tropical Medicine and Hygiene 99, 686–691. Homan, W.L. and Mank, T.G. (2001) Human giardiasis: genotype linked differences in clinical symptomatology. International Journal for Parasitology 31, 822–826. Hopkins, R.M., Constantine, C.C., Groth, D.A., Wetherall, J.D., Reynoldson, J.A. and Thompson, R.C.A. (1999) PCR-based DNA fingerprinting of Giardia duodenalis isolates using the intergenic rDNA spacer. Parasitology 118, 531–539. Inpankaew, T., Traub, R., Thompson, R.C.A. and Sukthana, Y. (2007) Canine parasitic zoonoses and temple communities in Thailand. Southeast Asian Journal of Tropical Medicine and Public Health 38, 247–255. Kettlewell, J.S., Bettiol, S.S., Davies, N., Milstein, T. and Goldsmid, J.M. (1998) Epidemiology of giardiasis in Tasmania: a potential risk to residents and visitors. Journal of Travel Medicine 5, 127–130.

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R.C.A. Thompson Kutz, S.J., Thompson, R.C.A., Kandola, K., Nagy, J., Wielinga, C., Polley, L. and Elkin, B. (2008) Giardia assemblage A: human genotype in muskoxen in the Canadian Arctic. Parasites and Vectors 1, 32 pp. Leonhard, S., Pfister, K., Beelitz, P., Wielinga, C. and Thompson, R.C.A. (2007) The molecular characterisation of Giardia from dogs in Southern Germany. Veterinary Parasitology 150, 33–38. Meloni, B.P., Lymbery, A.J. and Thompson, R.C.A. (1995) Genetic characterization of isolates of Giardia duodenalis by enzyme electrophoresis: implications for reproductive biology, population structure, taxonomy, and epidemiology. Journal of Parasitology 81, 368–383. Meyer, E.A. (1994) Giardia as an organism. In: Thompson, R.C.A., Reynoldson., J.A. and Lymbery, A.J. (eds) Giardia: From Molecules to Disease. CAB International, Wallingford, UK, pp. 3–15. Morrison, H.G., McArthur, A.G., Gillin, F.D., Aley, S.B., Adam, R.D., Olsen, G.J., Best, A.A., Cande, W.Z., Chen, F., Cipriano, M.J., Davids, B.J., Dawson, S.C., Elmendorf, H.G., Hehl, A.B., Holder, M.E., Huse, S.M., Kim, U.U., Lasek-Nesselquist, E., Manning, G., Nigam, A., Nixon, J.E. J., Palm, D., Passamaneck, N.E., Prabhu, A., Reich, C.I., Reiner, D.S., Samuelson, J., Svard, S. and Sogin, M.L. (2007) Genomic minimalism in the early diverging intestinal parasite Giardia lamblia. Science 317, 1921–1926. Nash, T.E., Herrington, D.A., Losonsky, G.A. and Levine, M.M. (1987) Experimental human infections with Giardia lamblia. Journal of Infectious Diseases 156, 974–984. O’Handley, R.M. and Olson, M.E. (2006) Giardiasis and cryptosporidiosis in ruminants. Veterinary Clinics of North America: Food Animal Practice 22, 623–643. O’Handley, R.M., Cockwill, C., McAllister, T.A., Jelinski, M., Morck, D.W. and Olson, M.E. (1999) Duration of naturally acquired giardiosis and cryptosporidiosis in dairy calves and their association with diarrhea. Journal of the American Veterinary Medical Association 214, 391–396. Olson, M.E., McAllister, T.A., Deselliers, L., Morck, D.W., Cheng, K.J., Buret, A.G. and Ceri, H. (1995) Effects of giardiasis on production in a domestic ruminant (lamb) model. American Journal of Veterinary Research 56, 1470–1474. Olson, M.E., O’Handley, R.M., Ralston, B.J., McAllister, T.A. and Thompson, R.C.A. (2004) Update on Cryptosporidium and Giardia infections in cattle. Trends in Parasitology 20, 185–191. Ramesh, M.A., Malik, S.B. and Logsdon, J.M., Jr (2005) A phylogenomic inventory of meiotic genes: evidence for sex in Giardia and an early eukaryotic origin of meiosis. Current Biology 15, 185–191. Read, C., Walters, J., Robertson, I.D. and Thompson, R.C.A. (2002) Correlation between genotype of Giardia duodenalis and diarrhoea. International Journal for Parasitology 32, 229–231. Rendtorff, R.C. (1954) The experimental transmission of human intestinal protozoan parasites. II. Giardia lamblia cysts given in capsules. American Journal of Hygiene 59, 209–220. Reynoldson, J.A., Behnke, J.M., Gracey, M., Horton, R.J., Spargo, R., Hopkins, R.M., Constantine, C.C., Gilbert, F., Stead, C., Hobbs, R.P. and Thompson, R.C.A. (1998) Efficacy of albendazole against Giardia and hookworm in a remote Aboriginal community in the north of Western Australia. Acta Tropica 71, 27–44. Robertson, I.D., Irwin, P.J., Lymbery, A.J. and Thompson, R.C.A. (2000) The role of companion animals in the emergence of parasitic zoonoses. International Journal for Parasitology 30, 1369–1377. Rodriguez-Hernandez, J., Canut-Blasco, A. and Martin-Sanchez, A.M. (1996) Seasonal prevalences of Cryptosporidium and Giardia infections in children attending day

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care centres in Salamanca (Spain) studied for a period of 15 months. European Journal of Epidemiology 12, 291–295. Sahagún, J., Clavel, A., Goñi, P., Seral, C., Llorente, M.T., Castillo, F.J., Capilla, S., Arias, A. and Gómez-Lus, R. (2007) Correlation between the presence of symptoms and the Giardia duodenalis genotype. European Journal of Clinical Microbiology and Infectious Diseases 27, 81–83. Savioli, L., Smith, H. and Thompson, A. (2006) Giardia and Cryptosporidium join the ‘Neglected Diseases Initiative’. Trends in Parasitology 22, 203–208. Sleeman, J.M., Meader, L.L., Mudakikwa, A.B., Foster, J.W. and Patton, S. (2000) Gastrointestinal parasites of mountain gorillas (Gorilla gorilla beringei) in the Parc National des Volcans, Rwanda. Journal of Zoo and Wildlife Medicine 31, 322–328. Smith, H.V., Cacciò, S.M., Tait, A., McLauchlin, J. and Thompson, R.C.A. (2006) Tools for investigating the environmental transmission of Cryptosporidium and Giardia infections in humans. Trends in Parasitology 22, 160–167. Steuart, R.F.L., O’Handley, R., Lipscombe, R.J. and Thompson, R.C.A. (2008) Alpha 2 Giardin is an assemblage A specific protein of human infective Giardia duodenalsis. Parasitology (in press). Tait, A. (1983) Sexual processes in the Kinetoplastida. Parasitology 86, 29–57. Teodorovic, S., Braverman, J.M. and Elmendorf, H.G. (2007) Unusually low levels of genetic variation among Giardia Lamblia isolates. Eukaryotic Cell 6, 1421–1430. Thompson, R.C.A. (2000) Giardiasis as a re-emerging infectious disease and its zoonotic potential. International Journal for Parasitology 30, 1259–1267. Thompson, R.C.A. (2004) The zoonotic significance and molecular epidemiology of Giardia and giardiasis. Veterinary Parasitology 126, 15–35. Thompson, R.C.A. (2008) Giardiasis: modern concepts in control and management. Annales Nestlé 66, 23–29. Thompson, R.C.A. and Monis, P.T. (2004) Variation in Giardia: implications for taxonomy and epidemiology. Advances in Parasitology 58, 69–137. Thompson, R.C.A., Reynoldson, J.A. and Mendis, A.H.W. (1993) Giardia and giardiasis. Advances in Parasitology 32, 71–160. Thompson, R.C.A., Reynoldson, J.A., Garrow, S.J., McCarthy, J.S. and Behnke J.M. (2001) Towards the eradication of hookworm in an isolated Australian community. Lancet 357, 770–771. Thompson, R.C.A., Traub, R.J. and Parameswaran, N. (2007) Molecular epidemiology of foodborne parasitic zoonoses. In: Murrell, K.D. and Fried, B (eds) Food-Borne Parasitic Zoonoses: Fish and Plant-Borne Parasites. Springer, New York, pp. 383–415. Traub, R.J., Monis, P.T., Robertson, I., Irwin, P., Mencke, N. and Thompson, R.C.A. (2004) Epidemiological and molecular evidence supports the zoonotic transmission of Giardia among humans and dogs living in the same community. Parasitology 128, 253–262. Traub, R.J., Monis, P.T. and Robertson, I.D. (2005) Molecular epidemiology: a multidisciplinary approach to understanding parasitic zoonoses. International Journal for Parasitology 35, 1295–1308. Troeger, H., Epple, H.-J., Schneider, T., Wahnschaffe, U., Ullrich, R., Burchard, G.-D., Jelinek, T., Zeitz, M., Fromm, M. and Schulzke, J.-D. (2007) Effect of chronic Giardia lamblia infection on epithelial transport and barrier function in human duodenum. Gut 56, 316–317. WHO (1996) Fighting Disease, Fostering Development: The World Health Report 1996. World Health Organization, Geneva.

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Cryptosporidium in Cattle: From Observing to Understanding R. FAYER, M. SANTÍN AND J.M. TROUT United States Department of Agriculture, Beltsville, MD, USA

Abstract Cryptosporidium parvum is a zoonotic pathogen transmissible from a variety of animals to humans and is a considerable public health concern. Dairy cattle have been identified in numerous reports as a major source of environmental contamination with this pathogen. However, the vast majority of these reports have been based on microscopic examination of the organism in faeces from cattle. This chapter traces the progress of research on bovine cryptosporidiosis from the first observations of infection to the present understanding of susceptibility on the part of the bovine host and pathogenicity on the part of the parasites that constitute the taxa under the umbrella of the genus Cryptosporidium. It includes information based on molecular typing with the SSU rRNA gene and subtyping with the GP60 gene, which enables epidemiologists and others to trace the sources of Cryptosporidium-related outbreaks.

Introduction Recent archaeological evidence from Egypt suggests that cattle were herded in prehistoric times, possibly from around 12,500 BC. It is not clear whether these cattle originally came from the ‘Fertile Crescent’. Initially, it is presumed, cattle were killed for food, later they were used as draught animals, and for thousands of years cows’ milk was consumed only by calves. As farming methods evolved, cattle became a major source of protein, milk and milk products, and leather. Over time, livestock production intensified to support growing human populations. Larger and larger herds became concentrated in increasingly smaller areas to maximize the efficiency of production. In industrialized countries, as well as less affluent areas of the world, the consequence of this evolution in animal husbandry was seen through the impact of enteric diseases on neonates, of which cryptosporidiosis is but one. Through the millennia of this close association between humans and cattle, enteric pathogens had billions upon billions of opportunities to adapt and to be readily transmitted between these hosts. Those that have are 12

© CAB International 2009. Giardia and Cryptosporidium: From Molecules to Disease (eds G. Ortega-Pierres et al.)

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of concern to public health and among them is the species Cryptosporidium parvum. Cryptosporidium infections have been reported in cattle worldwide. Before the development and application of molecular methods to aid in species determination, numerous publications simply documented the presence of Cryptosporidium oocysts in cattle faeces. Subsequently, many other publications have identified Cryptosporidium species in cattle but with limited information on prevalence and herd management. Cryptosporidiosis, especially in calves, has been associated with a wide range of clinical signs from no apparent ill effects to severe morbidity, resulting in poor performance and production losses and in some instances mortality. From the perspective of human health, cattle have often been implicated as a source of zoonotic Cryptosporidium species. Risk of human infection has been based on physical contact with cattle, contamination of fresh fruits and vegetables with manure, and manure runoff from farms into drinking water supplies. With the goal of producing healthy cattle while protecting food and water supplies, studies have been undertaken to obtain a clear understanding of the species of Cryptosporidium that infect cattle, the prevalence of infection, and the relationship of these species to the age of the animals. There has been much progress in defining the Cryptosporidium species, genotypes, and some subtypes present in cattle. This chapter traces the progress of research on bovine cryptosporidiosis from the first observations of infection in cattle to our present understanding of susceptibility and pathogenicity. It includes those parasites that constitute the taxa under the umbrella of the genus Cryptosporidium. Application of this information can benefit both animal and human health.

Early Reports of Cryptosporidiosis in Cattle The first report of bovine cryptosporidiosis appeared in 1971 and described stages of the parasite in tissue sections of the jejunum from an 8-month-old heifer with chronic diarrhoea (Panciera et al., 1971). This observation was soon followed by others in which diarrhoeic dairy and beef calves, 2 weeks old and younger, had similar infections (Barker and Carbonell, 1974; Meuten et al., 1974; Morin et al., 1976). The association between the parasite and illness became stronger when it was reported that cryptosporidia were probably common enteropathogens of calves (Pohlenz et al., 1978) and that Cryptosporidium was a pathogen in experimentally infected calves (Tzipori et al., 1983). A survey of neonatal dairy calves in Maryland, USA, found Cryptosporidium in healthy as well as in diarrhoeic calves (Leek and Fayer, 1984). This finding led to a study of factors contributing to clinical illness in calves experimentally infected with Cryptosporidium obtained from pooled calves’ faeces (Fayer et al., 1985). In that study, clinical illness was not consistently found in neonatal dairy calves experimentally infected with 3.2–30 × 106 oocysts, except when rotavirus and/or Clostridium perfringens was also present. It appeared that Cryptosporidium required the presence of other pathogens to produce illness. These findings gave rise to the following questions: 1. What was different in this study from reports in which cryptosporidiosis in calves was strongly associated with morbidity and mortality?

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2. Were other isolates of the parasite more pathogenic or did other studies simply lack viral and bacterial assessments? 3. Why were some naturally infected calves healthy and others severely ill? At that time there appeared to be a single species of Cryptosporidium that lacked host specificity, and answers to those questions would, in part, require the development of better taxonomic data. In the next few years, cattle were recognized as hosts for two species, C. parvum and C. muris (Anderson, 1987), infecting the intestine and the abomasum, respectively, and represented by small and large oocysts (C. parvum, ~4.5 × 5.5 mm and C. muris, ~5.5 × 7.4 mm). Molecular techniques and cross-species transmission studies eventually identified the abomasal form with the large oocysts as a new species, C. andersoni (Lindsay et al., 2000). Cryptosporidium parvum was then associated with diarrhoea in young calves and C. andersoni with asymptomatic adults. As reports accumulated it became apparent that under farm and field conditions calves acquired infections shortly after birth. The highest prevalence of cryptosporidiosis in cattle was found at 1–3 weeks of age, with oocysts excreted for an average of 12 days and diarrhoea, when present, lasting an average of 8 days (reviewed by Santín and Trout, 2008).

What is the Immune Status of Calves and How Does it Affect Susceptibility to Infection? Cell-mediated immunity had been shown to play an important role in control of C. parvum infections in mouse models, and by the early 1990s it was clear that HIV AIDS patients with low levels of T cells were extremely susceptible to cryptosporidiosis and other opportunistic pathogens, although little was known regarding the immune status of calves. To determine why young calves were so susceptible to infection, studies were designed to identify lymphoid cell populations at the site of infection in the ileum where lymphocytes could respond directly to the parasite (Pasquali et al., 1997; Canals et al., 1998). Intra-epithelial lymphocytes (IEL), and lamina propria lymphocytes (LPL) were collected from C. parvum-infected calves and uninfected control calves; the infected group was inoculated with oocysts at 1–2 days of age. Cells were collected from both groups at 7–9 days of age and analysed for phenotype and cytokine mRNA production. Significant increases in CD2+, CD3+, CD4+ and CD8+ T cells were observed in the IELs of infected versus uninfected calves. These findings were supported by reports of elevated IEL CD8+ T cells in infected versus uninfected calves (Wyatt et al., 1997) and of elevated LPL CD4+, CD8+ T, and g/d T cells in infected versus uninfected calves (Abrahamsen et al., 1997). IELs and LPLs from uninfected calves contained much lower percentages of CD4+ and CD8+ T cells than found in adult cattle (Pasquali et al., 1997) which could explain the age-related susceptibility of neonatal calves while supporting early observations of others that T cell subsets protect against cryptosporidiosis in mice (Ungar et al., 1991; Chen et al., 1993; McDonald et al., 1994) and humans (Flanigan et al., 1992). Correlated with

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the increases in CD4+ and CD8+ IELs was the finding of elevated IFN-g and IL-12 mRNA in IELs and LPLs from the ileum of infected calves. However, no significant increases were detected in mRNA levels for IL-2, IL-4 or IL-10 (Canals et al., 1998). These cytokine and phenotypic cell differences in primary infected versus uninfected neonatal calves indicated that the susceptibility of these young calves to enteric infections had an immunological basis, the absence or extremely low levels of T cells.

What Accounts for the Variability in Severity of Infections? Although pre-weaned calves are highly susceptible, infection with different isolates of C. parvum appears to result in variation in the severity of diarrhoea and number of oocysts produced. Such variation with different isolates has been demonstrated in human infectivity studies and in calves (Pozio et al., 1992; Okhuysen et al., 1999). Clinical cryptosporidiosis could not always be induced in experimental calf infections at inoculating doses of 3.2–30 × 106 C. parvum oocysts per calf (Fayer et al., 1985). Subsequently, inoculation of calves with 1 × 104 oocysts of a bovine C. parvum isolate from Alabama, USA, consistently induced severe clinical disease in experimentally infected calves (Fayer and Ungar, 1986). Another isolate of C. parvum from Auburn, Alabama (AUCP-1) was transmitted from one Cryptosporidium-naïve calf to another (26 calves) over a period of 3 years; these infections consistently resulted in oocyst production rates of 0.3–41.5 × 106/g of faeces for 1 day or more during the patent period, accompanied by profuse diarrhoea (Fayer et al., 1997). Then, unexpectedly, over a period of the next six serial passages through calves, the severity of clinical signs and oocyst output steadily decreased until inoculation with 1.5 × 106 oocysts resulted in no diarrhoea and the recovery of fewer than 1 × 106 oocysts per calf. A similar decrease in productivity and pathogenicity with the same isolate occurred at Colorado State University. The same isolate was also serially transmitted, over a 3-year period, through 40 groups of C57BL/6 mice (four mice per group) immunosuppressed with dexamethasone. The mice showed no clinical signs but oocyst output remained consistently high. When oocysts from the mice were transmitted to a calf, the AUCP-1 isolate once again exhibited the earlier pathogenicity and oocyst production. It was not determined what factors led to the loss of pathogenicity in progressive serial passage through calves or why pathogenicity was unaffected by passage through mice; however, one might speculate that the original isolate contained multiple genetically distinct subpopulations that were selected for or against during passage through a particular host.

Selection In an attempt to use selection to obtain a non-pathogenic precocious strain of C. parvum, an experiment was designed in much the same way as the two live commercial vaccines Paracox and Livacox were selected for in poultry (Fayer, 1994). Ten calves were serially infected with oocysts collected from the previous

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calf on the first day of the patent period. Instead of the expected decrease in pathogenicity, as seen with coccidia, the duration and severity of diarrhoea steadily increased from calf 1 through to calf 9, although no pattern was seen in number of oocysts shed per calf. Perhaps it was the ability of C. parvum to autoinfect, unlike the Eimeria species in the coccidiosis vaccines, that invalidated the concept of a precocious non-pathogenic immunizing strain of C. parvum. These observations of different isolates and of serial passage of specific isolates of C. parvum demonstrate that not only do differences in pathogenicity exist among field isolates but also that both the pathogenicity and fecundity of an isolate can change over time.

Double-stranded RNA Virus-like Particles Double-stranded RNA (dsRNA) virus-like particles have been found in Babesia, Trichomonas, Giardia, Leishmania and Eimeria (Hotzel et al., 1995). Two sizes of extrachromosomal dsRNAs were found in the cytoplasm of sporozoites of C. parvum and C. hominis but not in seven other species of Cryptosporidium (Khramtsov et al., 1997; Khramtsov and Upton, 2000). Sequence analysis showed distinctly different dsRNA sequences in isolates (species) from calves versus those from humans (Khramtsov et al., 2000). Small dsRNA sequences of isolates from 23 calves and 38 humans (Xiao et al., 2001) showed that isolates from the same outbreak had identical sequences; eight distinct nucleotide sequences were from cattle (C. parvum) and ten from humans (C. hominis). If any dsRNAs in Cryptosporidium are associated with pathogenicity this has not been demonstrated, but a recent study with our colleagues (M. Jenkins and J. Higgins, USDA, personal communication) has shown an association with fecundity. Calves infected with C. parvum-Beltsville oocysts produced substantially more oocysts than calves infected with C. parvum-Iowa oocysts. Increased fecundity was correlated with levels of C. parvum virus (CPV) as measured by real-time RT-PCR using CPV RNA-specific primers. The CPV signal in C. parvum-Beltsville sporozoites relative to C. parvum-Iowa was 3–4 times greater as measured by RT-PCR. The greater fluorescence intensity of C. parvum-Beltsville sporozoites labelled with antibodies to CPV 40 kDa capsid protein supported this observation. These findings suggest that CPV affects fecundity, which might in turn affect the severity of infection. What other factors affect severity of infection is not known.

Prevalence of Severe Morbidity and Mortality There is no formal reporting system for cryptosporidiosis in cattle. Reports of cases, surveys and outbreaks provide information, but not on the same basis as the public health reports of ProMed, FoodNet and MMWR. At the US Department of Agriculture in Beltsville, Maryland, telephone calls from farmers and veterinarians attributing high morbidity and mortality to cryptosporidiosis in dairy and beef calves were frequent, peaking in the late 1980s through to the

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early 1990s, but steadily decreased in number until in 2006 no such calls were received. Whether the actual number of severe outbreaks declined or whether the association of severe diarrhoea and death of calves with cryptosporidiosis was no longer a ‘new’ phenomenon, and therefore consultation was no longer sought, is not known. Nevertheless, this informal barometer of activity suggests a change in the frequency of severe and economically important outbreaks perhaps due to a time when an exceptionally pathogenic strain of C. parvum was present.

Species Other than C. parvum Cryptosporidium andersoni, which colonizes the gastric glands of the abomasum, has been reviewed by Santín and Trout (2008). Infections, primarily in calves older than 4 weeks of age, produce no diarrhoea or visible clinical signs. Infections are more chronic (sometimes lasting over a year) and oocyst production is less than that of C. parvum. Elevated plasma pepsinogen and decreased milk production have been attributed to infection. For other species and genotypes infecting cattle (Table 2.1), there are no reports of clinical signs, histological data or subclinical pathology.

Molecular Identification of Species in Cattle For decades, microscopy was the sole method for detecting oocysts, first by direct faecal smears routinely stained, and later stained by IFA techniques. Microscopy Table 2.1. Species and genotypes of Cryptosporidium found in cattle and locations where they have been detected (adapted from Santín and Trout, 2008). Cryptosporidium species/genotypes C. parvum C. bovis C. andersoni C. ryanae C. hominis C. suis C. suis-like Pig genotype II C. felis C. canis

Prevalence

Geographical location

Frequent Frequent Frequent Frequent Rare Rare (2 calves) Rare (3 cattle) Rare (1 cow) Rare (1 cow) Experimental infection only

Worldwide Worldwide Worldwide Worldwide Scotland, India, Korea USA and Zambia Denmark Denmark Poland USA

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was time-consuming and required highly trained personnel. Unless oocysts differed significantly in size, species identification was impossible. PCR was found to be more sensitive than IFA and, when combined with gene sequencing, could differentiate species and genotypes of Cryptosporidium. The most frequently used gene for identification has been the SSU rRNA (small subunit ribosomal RNA) gene (the 18S gene). Molecular analysis of Cryptosporidium from cattle has identified seven species and three genotypes: C. parvum, C. andersoni, C. bovis, C. canis, C. felis, C. hominis, C. suis, C. suis-like genotype, C. ryanae, and Cryptosporidium pig genotype II (Bornay-Llinares et al., 1999; Fayer et al., 2001, 2006; Santín et al., 2004; Smith et al., 2005; Geurden et al., 2006; Park et al., 2006; Starkey et al., 2006; Feng et al., 2007; Langkjær et al., 2007) (Table 2.1).

Prevalence of Species in Cattle In large-scale studies of cattle, C. parvum, C. andersoni, C. bovis and C. ryanae were found most frequently (Santín et al., 2004; Xiao et al., 2004; Fayer et al., 2006; Geurden et al., 2006; Feng et al., 2007; Langkjær et al., 2007). Other species and genotypes were found infrequently or rarely. Crytosporidium hominis was found in a few cattle in Scotland, India and Korea (Smith et al., 2005; Park et al., 2006; Feng et al., 2007), C. suis was found in one calf in the USA and another in Zambia (Fayer et al., 2006; Geurden et al., 2006), C. suis-like genotype and Cryptosporidium pig genotype II were found in three cattle and one cow in Denmark, respectively (Langkjær et al., 2007), C. felis was found in a cow in Poland (Bornay-Llinares et al., 1999), and C. canis infected calves experimentally but natural infections have not been detected (Fayer et al., 2001) (Table 2.1).

Large-scale Cross-sectional Study and Longitudinal Study of Species Prevalence Related to Age of Cattle A large-scale study involving 1615 cattle was conducted over a period of 4 consecutive years (Santín et al., 2004; Fayer et al., 2006, 2007). Each year 15 commercial dairy farms were included, two or three from each of seven states, ranging from Vermont to Florida along the east coast of the USA. Each year an attempt was made to collect faeces from 30 or more cattle from each farm. During the first 2 years, calves from 5 days to 11 months of age were examined. During the third year, dairy heifers 1–2 years of age were examined. During the fourth year, cows over 2 years of age were examined. Most of the farms visited in the first year were visited again in the following years, but in a few cases, because of management practices, either the required age or the required number of cattle were not available and substitute farms were introduced into the study. Based on SSU rRNA gene sequencing from PCR-positive specimens, virtually all infections in pre-weaned calves (<8 weeks of age) were C. parvum, with low levels of C. bovis, C. ryanae and C. andersoni in some calves. The number

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of C. parvum infections decreased markedly in post-weaned calves (2–11 months of age), while the number of calves with C. bovis, C. ryanae and C. andersoni increased; C. bovis was the predominant species in this age group. In cattle older than 12 months, C. andersoni was the most prevalent species. In Denmark, a similar pattern was observed: C. parvum was the predominant species in calves younger than 1 month of age, and C. bovis became the predominant species in calves 3–12 months of age (Langkjær et al., 2007). Both C. bovis and C. ryanae have been found to be geographically widespread in cattle (Santín et al., 2004; Fayer et al., 2006, 2007; Starkey et al., 2006; Feng et al., 2007; Langkjær et al., 2007). Cryptosporidium andersoni constitutes the majority of infections in mature cattle in Denmark, the Czech Republic, Japan and the USA (Enemark et al., 2002; Kvac and Vitovec, 2003; Sakai et al., 2003; Fayer et al., 2006, 2007; Kvac et al., 2006) with the exception that Langkjær et al. (2007) in Denmark, did not detect this species in cattle of any age. The prevalence of Cryptosporidium in cattle declines significantly with age (Santín et al., 2004; Fayer et al., 2006, 2007; Kvac et al., 2006; Langkjær et al., 2007). Therefore, species and genotypes found in mature cattle are actually present in relatively few animals. A longitudinal study involving 30 Holstein cattle on a single dairy farm was conducted over a 2-year period (M. Santín et al., 2008). Faeces were examined consecutively at weekly, two-weekly and then at monthly intervals from 1 week to 24 months of age for the presence of Cryptosporidium oocysts using a twostep nested PCR protocol to amplify the SSU rRNA gene. Every PCR-positive product was purified and sequenced. Cryptosporidium parvum, C. bovis, C. andersoni and C. ryanae were detected in every animal and their presence was strongly associated with the age of the animal. This longitudinal study strongly supports the findings of the cross-sectional studies. In the longitudinal study all pre-weaned calves were infected with C. parvum, followed in the post-weaned cattle by C. bovis and C. ryanae infections, and then in the older cattle C. andersoni infections.

Subtyping of C. parvum and C. hominis Because C. parvum is the zoonotic species most commonly identified in cattle, it became the focus of subtyping to better determine its genetic diversity and to provide a more accurate tool for source tracking. Most subtyping protocols have been based on the 60 kDa glycoprotein gene (GP60). Sequence analysis of GP60 has revealed several subtype families (alleles) and subtypes within those families for C. parvum and C. hominis (Cacciò et al., 2000; Strong et al., 2000; Peng et al., 2001, 2003; Alves et al., 2003; Tanriverdi et al., 2003; Wu et al., 2003; Xiao and Ryan, 2004). Most cattle isolates belong to the C. parvum subtype family IIa, which is the zoonotic subtype family. Within this family the most common subtype found worldwide was IIA15G2R1. For example, in Portugal 61 of 72 isolates were positive for this subtype (Alves et al., 2003, 2006). Likewise, in India 5 of 9 were positive (Feng et al., 2007), and in the USA 135 of 175 were positive for this subtype (Xiao et al., 2007). Greater diversity was found within allele IIa in

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Northern Ireland (Thompson et al., 2007). Subtypes IIaA18G3R1 and IIA15G2R1 were found in 120 and 28 of 214 isolates, respectively. The identity of IIA15G2R1 as the subtype most prevalent in calves and frequently detected in humans worldwide (Strong et al., 2000; Glaberman et al., 2002; Alves et al., 2003, 2006; Peng et al., 2003; Stantic-Pavlinic et al., 2003; Wu et al., 2003; Chalmers et al., 2005; Sulaiman et al., 2005; Abe et al., 2006; Trotz-Williams et al., 2006), as well as other subtypes identified by analysis of the GP60 gene, provides an important tool to accurately identify and track Cryptosporidium of veterinary and public health concern. The recent finding that C. parvum subtypes IIaA15G2R1 and IIaA17G2R1 were detected in stools from 12 laboratory-confirmed case patients who drank apple cider at a school outing in Ohio, and that subtype IIaA17G2R1 was also detected in the remains of some of the cider (Blackburn et al., 2006) illustrates the usefulness of subtyping. It is interesting to note that the same two subtypes were detected in a high percentage of faeces from cattle in Ohio (Xiao et al., 2007). Although these initial findings based on subtyping with GP60 have already served to provide a greater understanding of the transmission dynamics of C. parvum and C. hominis, the number of isolates examined is relatively few, so much more can still be learned. For other species and genotypes of Cryptosporidium affecting cattle and humans, additional typing tools are needed to reach a similar level of identification and understanding.

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Pohlenz, J., Moon, H.W., Cheville, N.F. and Bemrick, W.J. (1978) Cryptosporidiosis as a probable factor in neonatal diarrhea of calves. Journal of the American Veterinary Medical Association 172, 452–457. Pozio, E., Gomez Morales, M.A., Barbieri, F.M. and LaRosa, G. (1992) Cryptosporidium: different behaviour in calves of isolates of human origin. Transactions of the Royal Society of Tropical Medicine and Hygiene 86, 636–638. Sakai, H., Tsushima, Y., Nagasawa, H., Ducusin, R.J., Tanabe, S., Uzuka, Y. and Sarashina, T. (2003) Cryptosporidium infection of cattle in Tokachi District, Hokkaido. Journal of Veterinary Medical Science 65, 125–127. Santín, M. and Trout, J.M. (2008) Cryptosporidiosis of livestock. In: Fayer, R. and Xiao, L. (eds) Cryptosporidium and Cryptosporidiosis, 2nd edn. CRC Press, Boca Raton, FL, pp. 451–483. 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. Veterinary Parasitology 122, 103–117. Santín, M., Trout, J.M. and Fayer, R. (2008) A longitudinal study of cryptosporidiosis in dairy cattle from birth to 2 years of age. Veterinary Parasitology 155, 15–23. 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. Veterinary Record 156, 710–711. Stantic-Pavlinic, M., Xiao, L., Glaberman, S., Lal, A.A., Orazen, T., Rataj-Verglez, A., Logar, J. and Berce, I. (2003) Cryptosporidiosis associated with animal contacts. Wiener Klinische Wochenschrift 115, 125–127. Starkey, S.R., Zeigler, P.E., Wade, S.E., Schaaf, S.L. and Mohammed, H.O. (2006) Factors associated with shedding of Cryptosporidium parvum versus Cryptosporidium bovis among dairy cattle in New York State. Journal of the American Veterinary Medical Association 229, 1623–1626. 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. Infection and Immunity 68, 4117–4134. Sulaiman, I.M., Hira, P.R., Zhou, L., Al-Ali, F.A., Al-Shelahi, F.A., Shweiki, H.M., Iqbal, J., Khalid, N. and Xiao, L. (2005) Unique endemicity of cryptosporidiosis in children in Kuwait. Journal of Clinical Microbiology 43, 2805–2809. 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. Molecular and Biochemical Parasitology 130, 13–22. Thompson, H.P., Dooley, J.S.G., Kenny, J., McCoy, M., Lowery, C.J., Moore, J.E. and Xiao, L. (2007) Genotypes and subtypes of Cryptosporidium in neonatal calves in Northern Ireland. Parasitology Research 100, 619–624. 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. Parasitology Research 99, 346–352. Tzipori, S., Smith, M., Halpin, C., Angus, K.W., Sherwood, D. and Campbell, I. (1983) Experimental cryptosporidiosis in calves: clinical manifestations and pathological findings. Veterinary Record 112, 116–120. Ungar, B.L., Kao, T.C., Burris, J.A. and Finkelman, F.D. (1991) Cryptosporidium infection in an adult mouse model: independent roles for IFN-gamma and CD4+ T lymphocytes in protective immunity. Journal of Immunology 147, 1014–1022. Wu, Z., Nagano, I., Boonmars, T., Nakada, T. and Takahashi, Y. (2003) Intraspecies polymorphism of Cryptosporidium parvum revealed by PCR-restriction fragment length

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R. Fayer et al. polymorphism (RFLP) and RFLP-single-strand conformational polymorphism analyses. Applied and Environmental Microbiology 69, 4720–4726. Wyatt, C.R., Brackett, E.J., Perryman, L.E., Rice-Ficht, A.C., Brown, W.C. and O’Rourke, K.I. (1997) Activation of intestinal intraepithelial T lymphocytes in calves infected with Cryptosporidium parvum. Infection and Immunity 65, 185–190. Xiao, L. and Ryan, U.M. (2004) Cryptosporidiosis: an update in molecular epidemiology. Current Opinion in Infectious Diseases 17, 483–490. Xiao, L., Limor, J., Bern, C. and Lal, A.A. (2001) Tracking Cryptosporidium parvum by sequence analysis of small double-stranded RNA. Emerging Infectious Diseases 7, 141–145. Xiao, L., Fayer, R., Ryan, U. and Upton, S.J. (2004) Cryptosporidium taxonomy: recent advances and implications for public health. Clinical Microbiological Reviews 17, 72–97. Xiao, L., Zhou, L., Santín, M., Yang, W. and Fayer, R. (2007) Distribution of Cryptosporidium parvum subtypes in calves in eastern United States. Parasitology Research 100, 701–706.

3

Names Do Matter D.D. BOWMAN Cornell University, Ithaca, NY, USA

Abstract In zoology, each species is assigned a name by an expert or group of experts on the basis of characters that define an organism or groups of organisms that are designated as type specimens. As more is learned about a genus, the importance of certain characters for species designation may wax and wane, but the type materials are expected to remain constant and are usually held within museum collections for the purpose of later examination by additional experts as the need requires. Historically, in forms that have mating as part of the life cycle, the biological verification of a species has often been done, if possible, by crossing to prove that the offspring are viable and are capable of producing sustainable offspring themselves. Thus, in the case of Cryptosporidium species, crosses are potentially possible for species determination, but are not possible in the case of species within the genus Giardia. In both cases, however, it is expected that type specimens that correlate with the originally defined species will remain available for examination. Specific names sometimes have major impacts and ramifications that extend beyond the simple designations assigned to specimens by the experts naming a species. Thus, drug labels, if at all possible, contain the names of certain species of organisms, unless they can be shown to inactivate or kill more than one member of a species within a genus. Thus, a drug that kills the hookworm, Ancylostoma caninum, may have no effect on the related species, Ancylostoma braziliense; both names will not appear on the label unless the drug is shown to kill both species, and some drugs may kill only one species and not the other. It is possible that similar rationale could be applied to Cryptosporidium parvum in cattle; where a drug would have an effect on this species but would be without effect on the related Cryptosporidium bovis; this is typically what currently occurs with bovine coccidia. If a species of Giardia or Cryptosporidium is designated as zoonotic, regulations may be created narrowly for the control of one species and not another if the means exist for the ready designation of species. In the world of ecology and species protection, species or even subspecies are often protected against environmental encroachment or extinction, and this can have major implications for species protection by public and federal agencies. Names can be very important both scientifically and in the formation of policy.

Introduction The question as to the importance of names is not new. In 1920, Kofoid asked the same major questions regarding the taxonomy of Giardia (and also Cryptosporidium) that are still being asked today: © CAB International 2009. Giardia and Cryptosporidium: From Molecules to Disease (eds G. Ortega-Pierres et al.)

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D.D. Bowman The several species which have been described, especially by Benson (1908), are based on slight differences in form and proportions, and Noe (1908) raises the question as to the validity of these distinctions…We are inclined to regard Noe’s scepticism as not well grounded. This matter [of separate species] is one of more than restricted interest for it impinges on the one hand on the general biological question as to how far specificity of parasites to particular hosts obtains, and on the other hand on the very practical question as to whether or not rodents, especially mice, rats, and Belgian hares, are sources of infection to man, for these flagellate parasites, in mice and men at least, are often associated with chronic intestinal disturbances. (Kofoid, 1920)

These questions remain as the major concern of biologists today relative to these genera of organisms. Are different groups of organisms in different hosts restricted to those hosts? What is the zoonotic potential of the different groups of organisms relative to each other? Will the addition of names to the different species and assemblages bring clarity to the problem relative to the public and the medical community, or would they just serve to continue the confusion as to what to say when a diagnosis of cysts and oocysts is made? Would the clarification of species assist in the establishment of public policy for assessing the zoonotic risk of oocysts in drinking water or sewage effluents?

The International Code of Zoological Nomenclature The rules for names are set in the International Code of Zoological Nomenclature, published by the International Commission on Zoological Nomenclature (ICZN, 2006). This organization sets the rules for how names are made, defines what a taxon is and sets rules for how taxa are associated with a given name, decides what constitutes the publication of a name, and even has the power to suppress names for the purpose of maintaining stability. Because protozoa fall under the realm of eukaryotes in the world of zoology, they are covered by the code of zoological nomenclature rather than the codes that exist for fungi and bacteria. The code sets the rules for taxonomy from the names at the family level to the names used at the species level. The name of a species is a binomen (genus and species). The code also allows for the creation of subgenera (species aggregates within a genus) and subspecies within a species; this usage can actually form names consisting of three or four words. The use of species aggregates and subspecies may turn out to be very useful in the naming and categorization of organisms that appear on the many different cladograms and phenograms that have been created from the molecular identification of different clusters or taxa. Again, the code only specifically deals with the family through the species; higher taxonomic classifications are not bound by the code. The code also sets forth what the requirements are for describing a new species. Every new name published after 1999 must be explicitly indicated as intentionally new, the description must be associated with the explicit fixation of a holotype, or syntypes, for the nominal taxon and, where the holotype or syntypes are extant specimens, by a statement of intent that they will be (or have

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been) deposited in a collection along with a statement indicating the name and location of that collection. The code states the rules for publication, and paper publication is required, descriptions solely on the web are not recognized at this time. Authors are also cautioned that they should avoid confusing the literature by citing new names in abstracts or proceedings in a manner that does not really constitute publication, but will cause confusion as to priority. The code also establishes the principle of priority. The first name is the true name. However, it is possible that a name with priority may be discovered that has not been used for a very long time. This can be very confusing if another name is already entrenched in the literature. Thus, it is possible to suppress a name with priority under certain circumstances: Prevailing usage must be maintained when the following conditions are both met: the senior synonym or homonym has not been used as a valid name after 1899, and 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.

The goal here, as stated above, is the maintenance of stability amongst taxa. The standard of the code is the ‘type specimen’. The code goes to great lengths to describe different types and how to define a type: Each nominal taxon in the family, genus or species groups has actually or potentially a name-bearing type. The fixation of the name-bearing type of a nominal taxon provides the objective standard of reference for the application of the name it bears.

And: No matter how the boundaries of a taxonomic taxon may vary in the opinion of zoologists, the valid name of such a taxon is determined from the name-bearing type(s) considered to belong within those boundaries.

The neotype is an important concept describing species for which types may not be in existence or which have been destroyed. This may be used, for example, in the redescription of species of Giardia: Neotype is the name-bearing type of a nominal species-group taxon when no name-bearing type specimen (i.e. holotype, lectotype, syntype or prior neotype) is believed to be extant and an author considers that a name-bearing type is necessary to define the nominal taxon objectively.

And: A neotype is validly designated when there is an exceptional need and only when that need is stated expressly and when the designation is published with the following particulars: a statement that it is designated with the express purpose of clarifying the taxonomic status or the type locality of a nominal taxon; a statement of the characters that the author regards as differentiating from other taxa the nominal species-group taxon for which the neotype is designated; data and description sufficient to ensure recognition of the specimen designated; the author’s reasons for believing the name-bearing type specimen(s) (i.e. holotype, or lectotype, or all syntypes, or prior neotype) to be lost or destroyed, and the steps that

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D.D. Bowman had been taken to trace it or them; evidence that the neotype is consistent with what is known of the former name-bearing type from the original description and from other sources; however, a neotype may be based on a different sex or life stage, if necessary or desirable to secure stability of nomenclature; evidence that the neotype came as nearly as practicable from the original type locality and, where relevant, from the same geological horizon or host species as the original namebearing type; a statement that the neotype is, or immediately upon publication has become, the property of a recognized scientific or educational institution, cited by name, that maintains a research collection, with proper facilities for preserving name-bearing types, and that makes them accessible for study.

The code specifically states that the word ‘genotype’ should not be used in matters taxonomic. The reason is simply for clarity (from the Glossary of the Code: ‘Genotype: A term not recognized by the Code, formerly used for type species, but that should not now be used in zoological nomenclature’). The reasoning seems straightforward enough, and having access to words like ‘assemblage’ and ‘clade’, there probably are other highly useful alternatives to the use of the word ‘genotype’.

Ramifications In 1922, Hegner described Giardia canis very carefully according to the standards of the day. Also, he deposited type material: ‘The types and cotypes are deposited in the collections of the Department of Medical Zoology, School of Hygiene and Public Health, Johns Hopkins University’ (Hegner, 1922). The work on assemblages has shown that dogs may have their own Giardia in the form of assemblages C&D. So, what is to be done with Hegner’s description? Can the types be found? If so, what good are they for modern study? They were probably fixed with mercuric chloride and stained with haematoxylin after treatment with ferric ammonium sulphate; the chances seem good, therefore, that there will be no DNA available for molecular analysis. So will neotypes have to be selected? Will assemblages C&D turn into subspecies, or will the assemblages be recognized instead as subgenera? At the same time, for a redescription of Giardia canis and neotype selection, care would have to be taken to try and represent a collection of organisms similar to those that Hegner had in mind in his original description.

Splitters Splitters are currently holding sway in the taxonomy of Giardia and Cryptosporidium. The dissection of Cryptosporidium parvum has moved along relatively easily because there were almost no species described before the development of modern molecular methods. However, in the case of Giardia, there were many species described previously, so the creation of new names is much more complicated. This leaves open the question as to what is required to name a species, and instead of talking species, we discuss assemblages.

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Splitting can have major ramifications, especially in the field of regulatory affairs and liability. Classic cases include the environmental ramifications caused by the description of the snail darter, the spotted owl, and various subspecies of gorillas (Etnier, 1976; Noon and McKelvey, 1996; Jensen-Seaman et al., 2004). These have all had major fiscal, ecological and social implications for large geographical areas, with huge amounts being spent on litigation and management expenses. The same thing has happened with Giardia and Cryptosporidium. During the period when they were each considered to consist of a single species infecting all mammals other than rodents, the great fear was zoonosis, mainly fuelled by the devastating effects of cryptosporidiosis in AIDS patients. Giardia and Cryptosporidium received NIH funding, there were all sorts of funds spent for improved diagnostic methods for cysts and oocysts in drinking water, regulations were passed by the EPA with regard to allowable levels in water, and there was significant discussion by the EPA and USDA about the regulation of cysts and oocysts leaving confined animal feeding operations (CAFOs). Very large amounts of money and effort were expended. So, now the splitters are in charge. The discovery that the only major zoonotic species of Cryptosporidium in cattle is Cryptosporidium parvum, which is found only in young calves, means that it is going to be easier to manage the relatively small quantity of calf waste for pathogen control. Similarly, the recognition of Cryptosporidium canis and Cryptosporidium felis as parasites of dogs and cats, respectively, that only very, very rarely occur in people, can put pet owners’ and veterinarians’ minds at ease when dealing with a diagnosis in the family pet. Concerns about liability are real. If a pet is shedding Giardia cysts, is it zoonotic? Dogs without any clinical signs have been treated repeatedly with the goal of removing all cysts from their faeces. In some cases, this has resulted in very large bills for the client for an infection that did not seem to be bothering the dog and very probably was not putting the family at serious risk. The use of a separate name for the species found in the dog would make it easier for the veterinarian to tell the owners that they did not have to worry, and the veterinarian would not have to worry about the possibility of having to appear in court to defend his decision not to treat the asymptomatic animal.

Conclusion Names matter greatly, both for scientists and the public. Names should bring together information that adds to clarity as to what a species really is, should assist in understanding basic biology and assumed phylogenetic relationships, and allow discussion as to what risks a given species may pose to the health of humans and animals. The system does seem to be working in the realm of these two pathogens, and the taxonomy is sorting itself out. The work and methods being applied are giving us new insights on a regular basis. The problem is that the discoveries resulting in new taxonomy and systematics of the two different groups are being made too quickly for some and too slowly for others. However, it does appear that the community is learning and that it is working towards a sense of consensus that, of course, will probably never reach perfect agreement

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among gatherings of hundreds of very interested scientists intimately familiar with many different aspects of the organisms under discussion.

References Etnier, D.A. (1976) A new percid fish from the Little Tennessee River, Tennessee. Proceedings of the Biological Society of Washington 88, 469–488. Hegner, R.W. (1922) A comparative study of the Giardias living in man, rabbit, and dog. American Journal of Hygiene 2, 442–454. ICZN (2006) International Commission on Zoological Nomenclature. Available at http:// www.iczn.org/index.htm Jensen-Seaman, M.I., Sarmiento, E.E., Deinard, A.S. and Kidd, K.K. (2004) Nuclear integrations of mitochondrial DNA in gorillas. American Journal of Primatology 63, 139–147. Kofoid, C.A. (1920) A critical review of the nomenclature of human intestinal flagellates Cercomonas, Chilomastix, Trichomonas, Tetratrcihomonas, and Giardia. University of California Publications in Zoology 20, 145–168. Noon, B.R. and McKelvey, K.S. (1996) Management of the spotted owl: a case history of conservation biology. Annual Reviews in Ecology and Systematics 27, 135–162.

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Centenary of the Genus Cryptosporidium: From Morphological to Molecular Species Identification J. ŠLAPETA University of Sydney, NSW, Australia

Abstract The biology and species diversity of the genus Cryptosporidium is mystifying for many protozoologists and even more so for non-specialists. Historically, two morphologically distinct parasites of the gastrointestinal tract were originally described by Ernest E. Tyzzer from mice (i.e. Cryptosporidium muris and Cryptosporidium parvum). Later, these two parasites were thought to parasitize all mammals, including cattle. However, such a scheme became insupportable and human-to-human transmission is now associated with Cryptosporidium hominis. The causative parasite of the zoonotic animal-to-human transmission is Cryptosporidium pestis. In this chapter, all the species names applied to the genus Cryptosporidium are reviewed in the light of historical records and molecular taxonomy initiatives. A DNA approach to taxonomy stands on the implicit assumption that the reference databases used for comparison are sufficiently complete and feature-rich, with annotated entries. However, the uncertain taxonomic reliability of annotations in public DNA repositories forms a major obstacle to sequence-based species identification. Finally, but importantly, a huge gap exists between the number of described names and the number of identified genotypes. The closure of this gap represents a prime challenge for the decades to come.

Introduction Species of the genus Cryptosporidium have attracted a great deal of attention since the late 1970s as an object for epidemiological and molecular biological studies. However, while going through the published records, one is quickly confronted with the masked identity of the organism. There is a dramatic difference between the features of C. parvum as reported in 1980, 2000 and nowadays. The huge number of names and the jargon used to describe some isolates, along with the inability to continuously culture these parasites in vitro, has become a major obstacle to the comparison of experimental results from different laboratories over the century of existence of the genus Cryptosporidium, which was first described by Ernest E. Tyzzer (Fig. 4.1). © CAB International 2009. Giardia and Cryptosporidium: From Molecules to Disease (eds G. Ortega-Pierres et al.)

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Fig. 4.1. Portrait of Ernest E. Tyzzer, who described Cryptosporidium muris from a mouse stomach in 1907. Courtesy of the National Library of Medicine, negative no. 85-204.

Traditionally, taxonomy is based on morphology. However, for microbes such as Cryptosporidium species, morphological differences are minute or do not exist at all. This fact has been repeatedly emphasized, and additional characteristics, including site of infection and host specificity, proved to be useful for species identification. But while these biological characteristics were useful, they are largely impractical for routine identification. Thus, an accurate identification technique using molecular methods has been particularly welcome. Using molecular techniques, formerly neglected organisms have been rediscovered and others have become topical. The following is an account of advances in the taxonomy of the genus Cryptosporidium. The aim was to consider old reports based on morphology in the context of more recent work on DNA sequencing, which is generating ‘barcodes’ of these life forms. A consensus is presented within the rules of the International Commission of Zoological Nomenclature (1999) along with an annotated checklist of each species. Additionally, as part of the description, available DNA accession numbers of small subunit rDNA (SSU rDNA), Cryptosporidium oocyst wall protein 1 (COWP1), actin, and cytosolic 70 kDa heat shock protein (HSP70)

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gene sequences from reference specimens are tabulated as molecular identifiers of the respective species.

Mounting Complexity: the Value of Host Specificity Cryptosporidium was an obscure genus and very few reports were published prior to the 1980s. The genus Cryptosporidium Tyzzer 1907 was established for C. muris parasitizing mouse gastric glands (Tyzzer, 1907, 1910). A related sporozoan from the mouse intestine was named C. parvum (Tyzzer, 1912). Tyzzer correctly identified and experimentally verified the life cycle and, based on his histological findings, he correctly speculated about autoinfection within the host. It all seemed relatively simple, and common wisdom dictated that each host would have its own species. Providing the proof for this concept was very illfated. Vetterling et al. (1971) published a revision of the taxonomy and amended the description of the genus Cryptosporidium. Ironically, in contrast to Tyzzer’s original precise work, Vetterling et al. (1971) failed to identify oocysts as a source of infection. At the same time, Vetterling et al. (1971) described host-specific C. wrairi from the guinea pig. Unfortunately, C. wrairi has a peculiar nature, since it is a species never recovered from the host in the wild but only reported on a handful of occasions in guinea pig laboratory colonies. Nevertheless, host specificity was established as the sole character for species identification within the genus Cryptosporidium. Follow-up papers started to introduce new names haphazardly, solely on the basis of different host species. The life cycle and transmission were deemed elusive until the end of the 1970s (Bird and Smith, 1980) despite accumulating information on the significance of the parasite in immunocompromised hosts. Oocysts, the exogenous stage, were re-demonstrated as the source of the infection, and the complete life cycle was experimentally confirmed and validated on C. felis (Iseki, 1979). The genus description was amended to include the oocyst with four sporozoites in accordance with Tyzzer’s (1910) early description. Using the putative species host specificity, many new species were quickly described and named (i.e. Barker and Carbonell, 1974). Other evidence, however, suggested the existence of only a single species in the genus (Tzipori et al., 1980). Increasing medical interest due to the recognition of acquired immune deficiency syndrome (AIDS) caused an avalanche of publications describing numerous aspects of Cryptosporidium from the early 1980s onwards. Early studies supported the animal-to-human transmission of Cryptosporidium spp., but some data already suggested person-to-person transmission or human infections not acquired directly from infected animals (Casemore et al., 1985). Host specificity studies and morphological diagnostic techniques definitely delimitated the life cycle and identified oocysts as a primary cause of infection (Current and Reese, 1986; Fayer and Ungar, 1986). Experimental infections and ultrastructural studies provided evidence for the thin-walled, autoinfective oocysts and recycling type I meronts, thereby explaining overwhelming host infections after exposure to inocula containing small numbers of oocysts (Current and Reese, 1986).

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To simplify the mounting complexity of the taxonomy, Levine (1984) reviewed existing cross-transmission experiments and hosts of Cryptosporidium spp. and suggested the existence of only a single species in each of the animal groups: mammals, birds, reptiles and fish. This idea did not agree with the original description for the localization of gastric C. muris and intestinal C. parvum in mice (Tyzzer, 1912). A response to Levine’s paper was published by Upton and Current (1985), clearly identifying two morphologically distinct Cryptosporidium spp. in cattle. Both C. parvum and C. muris were deemed to be prototype species for all mammalian species until further evidence suggested otherwise (Upton and Current, 1985). Early isoenzyme analysis and later polymerase chain reaction (PCR)-based diagnostic techniques provided new evidence for the existence of human-tohuman, animal-to-human and waterborne transmission (Awad-el-Kariem et al., 1995; Morgan et al., 1995). Introduction of DNA sequencing provided the data to identify genetic variants affecting different or even the same host species. It seemed logical to believe that individual hosts each have their own unique hostspecific Cryptosporidium spp., as well as species that are less host-specific affecting a wider spectrum of hosts (Xiao et al., 2004). Molecular tools provided the techniques for accurate diagnosis and genesequencing-generated data which allowed the construction of molecular phylogenies mapping the evolutionary relationships between individual species and isolates. All this effort led to the reopening of the question: ‘Is the number of species currently recognized sufficient, or do we need to define more?’

Systematics and Initial Nomenclatural Considerations Systematics is a fundamental discipline in biology, consisting of three ideally inseparable processes: (i) the identification of a species with its characteristics and the description of a type specimen; (ii) applying taxonomy to name a species or taxon; and (iii) phylogenetics, which describes the relationships between species and other taxa. The major product of systematics is a classification system of species. The system, an anthropocentric view on the species history, is a flexible structure of arbitrary taxa. Understandably, there is controversy about the boundaries of each taxon, due to the existence of several opinions on what constitutes a species, i.e. species concepts. Before one starts to talk about species in the genus Cryptosporidium, it is important to distinguish between two distinct elementary concepts. 1. The species is a biological entity that is represented by a distinct population of individuals. The definition of a species is a controversial issue and many theories exist, each concept favouring some aspects over others (e.g. biological species concept, phylogenetic species concept, etc.). Importantly, the species is based on the preferences of the group of researchers and their own local judgement, experience and consensus applying the ‘whatever species concept’ in favour at the time. With no strict rules preset, the biologists in each field tend to be either those who prefer to group several possible species into a single species, thus

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representing the ‘lumpers’ category; or those who tend to divide existing species into new species (who are therefore called ‘splitters’). In the case of parasites, clinicians often seem to represent the first category, aligning with the conservative, less name-rich scenario, unlike zoologists, who constantly modify and adjust the systematic scheme. 2. The name is the binominal name of a species consisting of Latin genus and species names, and is an arbitrary but very useful identifier of the species. The name must be unique, to make communication easier between researchers. In contrast to the species definition, the rules for naming species are set very strictly within an international code. In the case of the genus Cryptosporidium, as well as for the majority of the parasitic protozoa, this is the International Code of Zoological Nomenclature (ICZN), specifically the 4th edition (International Commission on Zoological Nomenclature, 1999), which supersedes all previous editions as from 1 January 2000. It stands on the holotype, the name-bearing specimen of the species that the author has used for the original description. The holotype is deposited in a museum and is the reference material for the species. However, in the sense of protistan terminology, the term holotype (= type) is substituted with hapantotype, i.e. one or more preparations consisting of directly related individuals representing distinct stages of the life cycle (Article 72.5.4, 73.3). Importantly for Cryptosporidium spp., the type (= hapantotype) is the specimen or specimens illustrated and/or photographed or described, but not the illustration/photograph itself or its description (Article 72.5.6). Thus, in fact, for the majority of protozoa there is no original museum material for subsequent investigators to use for comparative purposes, apart from the original printed publication. On the other hand, the fact that the specimens illustrated no longer exist or cannot be traced does not invalidate the type designation (Article 73.1.4). The genus Cryptosporidium was traditionally classified within coccidia, where the oocysts contain, in general, sufficient morphological differences to allow identification. Such a model worked quite well for the scenario described by Upton and Current (1985) with respect to distinguishing between the stomach and the intestinal species in cattle. The morphological detail of the oocyst proved quite practical for species such as Eimeria, and high-quality micrographs of sporulated oocysts along with validated data were considered to form an acceptable substitute for the type specimens (Duszynski, 1999). However, taking into consideration the 4th edition of the ICZN (International Commission on Zoological Nomenclature, 1999), such a practice had two drawbacks: (i) illustration is not a syntype, i.e. name-bearing type (Article 72.5); and (ii) the type must be fixed (Article 72.3). Even so, Cryptosporidium oocysts possess very few morphological characteristics and consequently the identity of Cryptosporidium species cannot usually be assigned using oocyst morphology alone (Fall et al., 2003); therefore such illustrations are of limited diagnostic and descriptive value. Taking into account all of the above, all existing names in the genus Cryptosporidium were re-evaluated in accordance with the ICZN and categorized as follows: 1. The names are reported as nomen dubium (s.)/nomina dubia (pl.) for those species that are species inquirenda (s.)/species inquirendae (pl.), that are impossible

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to recognize and identify again but where the name was based on correct rules of the code. These species are placed to one side until a detailed review is initiated. 2. The names incorrectly defined, i.e. without respecting ICZN, are assigned as nomen nudum (s.)/nomina nuda (pl.). The name is unavailable, but upon new description can be made available for the same or a different concept. For species where there is evidence of an original misidentification of a different organism, the names and species are removed from Cryptosporidium. 3. Otherwise, the name is valid. So much for the technicalities of taxonomy; now let’s apply the above to the Cryptosporidium species names (Table 4.1). In total, throughout the 100 years that the genus Cryptosporidium has been in existence, 37 names have been introduced. Of these, four names are nomina nuda and five have been removed from the genus as being different organisms (e.g. Sarcocystis crotali – syn. Cryptosporidium crotali). The remaining 28 names are treated here as available names; nevertheless, five of these currently need redescription (i.e. C. anserinum, C. agni) or remain aside due to insufficient information (i.e. C. rhesi, C. garnhami). This leaves us with 23 names to be associated with individual species. Here, I recognize in total 21 species; thus two names are reported as synonyms (C. tyzzeri, C. saurophilum). On one hand, the majority of these have a dominant host, but on the other hand, they are accidentally found in possibly aberrant hosts; as is evidenced by rare findings of diverse Cryptosporidium isolates in human and calf faeces. The exception to this rule is the zoonotic C. pestis, which has the potential to temporarily affect the majority of mammalian hosts, and C. meleagridis, which is known to be the only species affecting birds as well as mammals (Table 4.1).

Nucleotide Sequence Data: Reference Material – ‘Barcode’ Approach Nucleotide data have played a dominant role in revealing the epidemiology and Cryptosporidium population structure. While Cryptosporidium isolates remain difficult to grow in vitro, the use of PCR and animal models have provided muchneeded characterization of many former morphologically defined species. Several single-locus analyses provided a secure foundation for multiple previously uncharacterized species. The leading role was given to SSU rDNA as the most universal marker, soon followed by additional conservative markers (actin, HSP70). Supportive evidence has also come from COWP1. Characterization of the sequence of these four genes from a diverse selection of isolates has provided the groundwork for follow-up studies that mapped new isolates within the existing diversity of each of these genes. These four markers are the universal and dominant markers used to characterize individual isolates, and the availability of existing sequences makes them a suitable entry point for any survey. Since the late 1990s there has been a steady increase in the number of individual sequences in public databases (Fig. 4.2). The SSU rDNA gene remains the preferred

Update on Cryptosporidium spp. names and species status.

Name

Available/ Unavailable

1 2 3 4 5 6 7

C. muris Tyzzer 1907 C. parvum Tyzzer 1912 C. crotali Triffit 1925 C. vulpis Wetzel 1938 C. baikalika Matschoulsky 1947 C. meleagridis Slavin 1955 C. tyzzeri Levine 1961

Available Available Removed* Removed* Removed* Available Available

8

Removed*

12

C. lampropeltis Anderson, Duszynski & Marquardt 1968 C. ameivae Arcay de Peraza & Bastardo de San Jose 1969 C. ctenosauris Duszynski 1969 C. wrairi Vetterling, Jervis, Merill & Sprinz 1971 C. agni Barker & Carbonell 1974

13

C. anserinum Proctor & Kemp 1974

Available

14

C. bovis Barker & Carbonell 1974

Available

15

C. cuniculus Inman & Takeuchi 1979

Available

NN

9 10 11

Host range NS

*

Homo

Valid – redescribed by Tyzzer (1910) Valid

1 2

M M

Yes

Valid Subjectively invalid (junior subjective synonym with C. meleagridis)

3

MB

Yes

4

M

5

M

6

M

Name status/Species status

Unavailable

Nomen nudum/incomplete description

Removed* Available

Valid

Available

Subjectively invalid (nomen dubium/species inquirenda) Subjectively invalid (nomen dubium/species inquirenda) Valid – redescribed by Fayer, Santín and Xiao (2005) Valid

Bos

Centenary of the Genus Cryptosporidium

Table 4.1.

Yes

37

(Continued)

38

Table 4.1.

Continued Host range

Name

16

C. felis Iseki 1979

Available

Valid

17

C. rhesi Levine 1980

Available

18 19

C. serpentis Levine 1980 C. garnhami Bird 1981

Available Available

20

Available Unavailable

Nomen nudum/incomplete description

Available

Valid

Available Unavailable

Subjectively invalid (nomen dubium/species inquirenda) Nomen nudum/incomplete description

Available

Valid

11

Available Available

Valid Valid

12 13

F F

28

C. nasoris Hoover, Hoerr, Carlton, Hinsman & Ferguson 1981 C. enteritidis Payne, Lancaster, Heinzman & McCutchan 1983 C. baileyi Current, Upton & Haynes 1986 C. curyi Ogassawara, Benassi, Larsson & Hagiwara 1986 C. villithecum Paperna, Landsberg & Ostrovska 1986 C. varanii Pavlásek, Lávicˇková, Horák, Král & Král 1995 C. cichlidis Paperna & Vilenkin 1996 C. reichenbachklinkei Paperna & Vilenkin 1996 C. saurophilum Koudela & Modrý 1998

Subjectively invalid (nomen dubium/species inquirenda) Valid – redescribed by Tilley et al. (1990) Subjectively invalid (nomen dubium/species inquirenda) Valid

Available

29

C. galli Pavlásek 1999

Subjectively invalid (junior subjective synonym with C. varanii) Valid – redescribed by Ryan et al. (2003)

14

B

21 22 23 24 25 26 27

Available

Name status/Species status

NS

*

Homo

Bos

7

M

Yes

Yes

8

R

9

F

10

B

J. Šlapeta

NN

Available/ Unavailable

31 32

33

34 35

36

37

C. andersoni Lindsay, Upton, Owens, Morgan, Mead & Blagburn 2000 C. canis Fayer, Trout, Xiao, Morgan, Lal & Dubey 2001 C. blagburni Morgan, Monis, Xiao, Limor, Sulaiman, Raidal, O’Donoghue, Gasser, Murray, Fayer, Blagburn, Lal & Thompson 2001 C. hominis Morgan-Ryan, Fall, Ward, Hijjawi, Sulaiman, Fayer, Thompson, Olson, Lal & Xiao 2002 C. molnari Alvarez-Pellitero and Sitja-Bobadilla 2002 C. suis Ryan, Monis, Enemark, Sulaiman, Samarasinghe, Read, Buddle, Robertson, Zhou, Thompson & Xiao 2004 C. scophthalmi Alvarez-Pellitero, Quiroga, Sitjà-Bobadilla, Redondo, Palenzuela, Padrós, Vázquez & Nieto 2004 C. pestis Šlapeta 2006

Available

Valid

15

M

Yes

Yes

Available

Valid

16

M

Yes

Unavailable

Nomen nudum/incomplete description

Available

Valid

17

M

Yes

Yes

Available

Valid

18

F

Available

Valid

19

M

Yes

Yes

Available

Valid

20

F

Available

Valid

21

M

Yes

Yes

Centenary of the Genus Cryptosporidium

30

* (from Cryptosporidium) Notes: NN – name number; NS – currently recognized species number; M, mammal; B, bird; R, reptile; F, fish.

39

40

J. Šlapeta

Number of sequences in nucleotide databases

500 small subunit rDNA oocyst wall protein 1 400

actin 70 kDa heat shock protein 60 kDa glycoprotein precursor

300

200

100

0 1997

1998

1999

2000

2001 2002 Year

2003

2004

2005

2006

Fig. 4.2. The increasing numbers of individual SSU rDNA, COWP1, actin, HSP70 and GP60 gene sequences of Cryptosporidium spp. in nucleotide databases over the 10 years from 1997 to 2006.

marker in recent studies, with over 430 available sequences to compare, followed by COWP1 with some 130 available sequences. The sequence numbers of actin and HSP70 are appreciably lower, due to difficulties with the amplification of these genes from some materials. While the above genes remain extremely useful in primary characterization of the individual isolates, especially from animal hosts, it soon became apparent that for fine epidemiological studies these markers are too conservative. A 60-kDa glycoprotein precursor (GP60) proved to be the ideal marker for the elucidation of the predominant human-derived isolates. Nowadays, besides SSU rDNA, human epidemiological studies require characterization of GP60, with the comparison of over 260 available sequences being possible (Fig. 4.2). These numbers underestimate the real number of genotypes made, due to the absence of published genotyping results in the nucleotide databases. The current practice of submitting only the unique sequences, while referencing the identity to known sequences should be avoided. Curators of the primary nucleotide databases (NCBI, EMBL, DDBJ) recommend the submission of complete datasets of all sequences regardless of the identity match with already available sequences. Such a practice, with thorough annotation, will lay the foundations for quantitative meta-analyses with direct reference to features associated with these sequences, i.e. host species. The available sequences in public databases are an extremely valuable source of information; nevertheless, they represent secondary material and thus, in general, their reliability may be questionable (Harris, 2003; Morrison, 2006).

Centenary of the Genus Cryptosporidium

41

As well as the quality of the nucleotides reported, the accuracy and the depth of annotation remains a serious issue (Morrison, 2006; Nilsson et al., 2006). First, individual submitters provide their preferred species name, which is then retained in the database. However, the definition of a species often differs between researchers and over time. Since the late 1990s, ten new names have been introduced (Table 4.1). For many of these species, DNA has been the primary source of evidence for their distinct status. First these individuals acquired type/isolate/ genotype status within known species or only as a Cryptosporidium sp. For example, C. canis sequence accession numbers may be found in three distinct organism ranks despite representing a single valid species; i.e. SSU rDNA is AF112576, but this accession number retains its definition as ‘Cryptosporidium parvum strain CPD1’, COWP1 is AF266274, defined as ‘Cryptosporidium sp. 715’, HSP70 AF221529 is defined as ‘Cryptosporidium parvum isolate 244’, and finally actin AF382340 is correctly defined as ‘Cryptosporidium canis isolate 715’. Second, the annotation of many sequences does not provide sufficient information to associate the sequence with its host, isolate number, country of origin, etc. The use of common names for hosts is insufficient, i.e. monkey or snake. It may be argued that such information appears in the paper; however, in many cases, this information is absent there as well. It is now imperative to take advantage of the ‘features’ entry currently available during the process of sequence submission including: ‘country’, ‘specific_host’, ‘isolate’ and ‘dev_stage’. Unfortunately, the popular genotype status is not a recognized feature, but generally it is acceptable to provide this information in the ‘note’ field. To reduce the ambiguity present in the database derived from a diverse array of submitters, I provide a table with reference sequence accession numbers (Table 4.2). This table aims to list all available full-length sequences, complete where possible, for each named species of the genus Cryptosporidium (Table 4.1). I have taken all precautions to choose the most representative and complete sequence, if multiple sequences exist in databases. These sequences can be viewed as the ‘barcodes’ of individual species; moreover, as more data become available they will be improved and extended. Where possible, I chose to use only single sequences, to avoid future misinterpretations resulting from the construction of a consensus sequence. One exception is the SSU rDNA of C. galli, which is a consensus of two overlapping partial sequences. The four genes SSU rDNA, COWP1, actin and HSP70 are the most commonly used markers in genotyping analyses and species delimitation studies. The hypervariability of the GP60 marker prohibits its use in species identification; moreover, it is only available for the study of dominant populations which infect humans. The 21 recognized species are relatively well represented within the four genes (Table 4.2). The most problematic species are the fish species (five species), with only the assumed C. molnari species represented by SSU rDNA and actin. The COWP1 sequence is missing for C. bovis, C. cuniculus and C. galli. The list and completeness of the sequences may seem very strong, but multiple sequence alignments reveal the hidden incompleteness of many sequences in public databases. The partial sequence representation is further reflected in Cryptosporidium surveys, where only a diagnostic hypervariable region (400–800 bp) is amplified and sequenced, i.e. the use of SSU rDNA primers targeting the hypervariable

42

Table 4.2.

Reference sequences for named Cryptosporidium spp. SSU rDNA

COWP1

actin

HSP70

C. andersoni C. baileyi C. bovis C. canis C. cichlidis C. cf. cuniculus C. felis C. galli C. hominis C. hominis draft genome TU502 strain&&

AF093496 L19068 AY741305 AF112576 n.a. AY120901 AF108862 AF316624+AY168847*** AF108865 chromosome 2: Chro.rrn013 (6112–7866) chromosome 2: Chro.rrn021 (334–2082) chromosome 2: Chro.rrn022 (21763–23515) chromosome 7: Chro.rrn016 (23535–25283) chromosome 8: Chro.rrn005 (599–1001) chromosome 8: Chro.rrn006 (1064–1516) AF112574

AF266262 AF266276 n.a. AF266274 n.a. n.a. AF266263 n.a. AF266265 chromosome 6: Chro.60244

AF382352 AF382346 AY741307 AF382340 n.a. AY120924 AF382347 AY163901 AF382337 chromosome 5: Chro.50056

AF221542 AJ310880 AY741306 AF221529 n.a. AY273775 AF221538 AY168849 AF221535 chromosome 2: Chro.20010

AF266266

AF382351

AF221537

AY524773 AB089284

n.a. AF161579

AY524772 AF382350

n.a. AF221543

C. meleagridis (syn. C. tyzzeri) C. cf. molnari C. muris

J. Šlapeta

Species (alphabetically)

n.a. AF112571 AF108864 chromosome 1: cgd1_5 (46–743) chromosome 2*: cgd2_? (200603–202360) chromosome 7: cgd7_5535 (1278301–1278425**) chromosome 7: cgd7_7 (1582–3330) chromosome 8: cgd8_5425 (1155551–1156729)

n.a. AF266268 BX538351.1:61269..66140 chromosome 6: cgd6_2090

n.a. AF382343 M86241# chromosome 5: cgd5_3160

n.a. AF221530 U11761## chromosome 2: cgd2_20

C. reichenbachklinkei C. scophthalmi C. serpentis C. suis C. varanii (syn. C. saurophilum) C. wrairi

n.a. n.a. AF151376 AF115377 AF112573

n.a. n.a. AF266275 AF266270 AF266277

n.a. n.a. AF382353 AF382344 AF382349

n.a. n.a. AF221541 AF221533 AF221540

AF115378

U35027+U35028

AF382348

AF221536

Centenary of the Genus Cryptosporidium

C. nasoris C. parvum C. pestis C. pestis draft genome IOWA strain&

Notes: n.a. – sequences are not available; SSU rDNA – small subunit ribosomal DNA, >1500 bp (complete or almost complete) is desirable for comparative purposes; COWP1 – oocyst wall protein with type I and type II cysteine-rich repeats, oocyst EB module wall protein (Templeton et al., 2004); HSP70 – 70 kDa heat shock protein, cytosolic form; & – draft genome sequence of C. pestis IOWA (Abrahamsen et al. 2004, AAEE01000000) has five SSU rDNA (some are partial); for the purpose of this table chromosome 7: cgd7_7 sequence serves as reference sequence; LeBlancq et al. (1997) identified the total of five copies and two slightly different types of rDNA units per genome of C. pestis KSU-1: referred to as Type A (AF015772) and Type B (AF308600); && – draft genome sequence of C. hominis TU502 (Xu et al. 2004, AAEL01000000) has six SSU rDNAs (some partial); for the purpose of this table chromosome 7: Chro.rrn016 and chromosome 2: Chro.rrn022 as reference sequences; * – on chromosome 2, cgd2_1375 (299599–301356) is currently erroneously annotated as SSU rDNA, a different non-annotated region on the same contig AAEE01000005 is the true SSU rDNA; ** – cgd8_5425 (1155551–1156706); here several additional nucleotides are added as of the contig; *** – the sequence is concatenated from two overlapping partial sequences; # – the sequence from chromosome 3 (Kim et al., 1992) but genome projects of C. pestis Iowa and C. hominis TU502 both identified this gene on chromosome 5 (Abrahamsen et al., 2004; Xu et al., 2004); ## – the sequence reported by Khramtsov et al. (1995); see also mRNA sequence U69698 – unpublished, GenBank.

43

small subunit rDNA C. pestis IOWA (1749 nt) C. hominis TU502 (1753 nt)

200

150

100

50

0 0

5 -2

0

40

70 60 50 40 30 20 10 0

actin C. pestis IOWA (1131 nt) C. hominis TU502 (1131 nt)

70 kDa heat shock protein C. pestis IOWA (2022 nt) C. hominis TU502 (2034 nt)

1 50

0

51

0 10

0

0

5 12

01

0 15

51

0

01

5 17

0

0 20

51

30 25 20 15 10 5 0 0

0

80

7 12 15 10 17 Sequence length category (nt)

0

5 -7

35

5 -2

oocyst wall protein 1 C. pestis IOWA (4872 nt) C. hominis TU502 (4863 nt)

0 0 0 0 0 0 0 0 0 25 -50 -75 100 125 150 175 200 500 1 01 1- 1- 1- 1- 1- 15 5 75 00 25 50 75 00 2 1 1 2 1 1 Sequence length category (nt)

0

0 -5

1 25

Number of sequences in the length category

J. Šlapeta

0

0

0 -5

1 25

1 50

5 -7

0

0

51

0 10

01

5 12

7 10 Sequence length category (nt)

Number of sequences in the length category

Number of sequences in the length category

Number of sequences in the length category

44

0-

40 35 30 25 20 15 10 5 0

0 0 0 0 0 0 0 0 0 25 -50 -75 100 125 150 175 200 225 1 01 1- 1- 1- 1- 1- 15 2 5 75 00 25 50 75 00 2 1 1 1 1 Sequence length category (nt)

0-

Fig. 4.3. Representation SSU rDNA, COWP1, actin and HSP70 sequences available in primary nucleotide databases (NCBI/EMBL, DDBJ). Individual sequences were categorized according to their length. The full length of individual genes from C. pestis and C. hominis draft genomes is given above the graph.

region by Johnson et al. (1995), CPB-DIAGF and CPB-DIAGR primers or COWP1 by Spano et al. (1997), and Cry-15 and Cry-9 primers (Fig. 4.3). Many of the sequence entries for HSP70 and COWP1 are short in sequence length, in contrast to actin, where many species are almost fully sequenced. Moreover, many of the currently recognized genotypes are difficult to examine further due to the generation of only partial sequences by investigators.

Centenary of the Genus Cryptosporidium

45

It should be stressed that every effort should be made to generate full-length sequences for already existing species, genotypes or new genotypes being described. These sequences represent a permanent record upon which an investigation for biological characters can be based. To avoid the confusion that arises from the rapidly expanding complexity of species names and number of sequences generated in different laboratories since the late 1990s, a website has been developed (‘Taxonomy of the genus Cryptosporidium’ at http://www.vetsci.usyd.edu.au/staff/JanSlapeta/). This website aims to provide a common gateway for analysing Cryptosporidium species and to standardize the terminology of individual isolates. Alignments of the reference sequences from the named species are provided, with further detailed annotation of individual species. Furthermore, currently under development is a webbased analysis (BLAST, multiple sequence alignment, tree building) as well as an in-house annotation database of sequences to streamline and standardize the process for the identification of Cryptosporidium isolates.

Will the Real Cryptosporidium parvum Please Stand Up? When reviewing the large amount of information on the human and bovine derived isolates, it became apparent that the mouse species, C. parvum Tyzzer 1912, described and illustrated by Tyzzer (1912), is different from the commonly encountered species affecting humans and cattle (Šlapeta, 2006). Ernest E. Tyzzer described the parasite from the intestine of tame mice in his Harvard laboratory (Tyzzer, 1912). The species was easily transmissible to mice and was present in great numbers in the intestines of affected mice. In the absence of the type material and with a lack of data about the host specificity of Tyzzer’s original material, we need to subjectively align the available data with the current information. Two studies provided noteworthy details on Cryptosporidium spp. in mice (Mus musculus). The first study, by Klesius et al. (1986), determined up to 30% incidence of cryptosporidiosis in mice at a calving site and susceptibility of calves to some of these oocysts from mice faeces. In the second study, Morgan et al. (1999) genotyped many samples of Cryptosporidium spp. from mice obtained worldwide and genetically characterized a specific ‘mouse genotype’ using several independent nuclear loci distinct from known genotypes. This ‘mouse genotype’ was genetically different from the ‘bovine’ and ‘human’ genotypes previously identified in Australia and Maryland, USA, and lumped under the umbrella name C. parvum (Xiao et al., 2004). Thus, the genotyping of mice isolates provided additional evidence for the existence of the ‘bovine genotype’ in wild mice (Morgan et al., 1999). Consequently one can speculate that it was this ‘bovine genotype’ that was infectious for calves in the study of Klesius et al. (1986). This ‘bovine genotype’ essentially infects only neonatal mice and only affects adult mice transiently (Korich et al., 2000), unlike authentic rodent C. parvum isolates that produce heavy infections in adult laboratory mice (Bednarska et al., 2003). No experimental calf infections were conducted using the true C. parvum ‘mouse genotype’. Despite this fact, cattle and human isolates of Cryptosporidium

46

J. Šlapeta

spp. have attracted considerable attention in recent years, but no ‘mouse genotype’ (C. parvum) has been identified (Xiao et al., 2002; Santín et al., 2004). Therefore, C. parvum (C. parvum ‘mouse genotype’) appears to be host-specific, because either there are no natural calf or human infections or they represent an extremely rare case. It is generally agreed that the species that Tyzzer used for his description is indeed the ‘mouse genotype’ (Šlapeta, 2007; Xiao et al., 2007). An urgent step towards the stability of historical information is to apply Tyzzer’s name to the species he used for the description and to use appropriate names for the two dominant human species. The above findings led to the recent taxonomic treatment and support for the identity of the ‘mouse genotype’ with the original description, type host, experimental transmission and illustrations of C. parvum (Šlapeta, 2006). Cryptosporidium parvum sensu Tyzzer, 1912 is a host-adapted species of Mus musculus with no documented capacity to infect either humans or domestic cattle.

Consensus, Reversibility and Universality Two species which can by supported by morphological characteristics and by localization and development in the host are C. muris and C. parvum. The larger C. muris is found in the stomach, whereas the intestine is the location of the smaller C. parvum. The status quo of only these two species affecting mammals is retained by Upton and Current (1985), and is suggested to be practical unless further evidence shows otherwise. However, a diverse spectrum of hosts and distinct DNA has provided further evidence of multiple lineages with different evolutionary history, and thus provides only preliminary evidence for a species. Older names synonymized under the umbrella names C. parvum and C. muris have been re-erected as full species (i.e. C. felis, C. wrairi). Those for which no name was thought to be available and whose identification was based on DNA sequencing have acquired ‘genotype’ status or have been named as new species. However, mixing the latter with the former has introduced inconsistency to what is C. parvum. Indeed, the name C. parvum has lost its purpose; it is no longer a unique identifier. Genotypes now seem to be much more useful descriptors, i.e. human genotype, mouse genotype, canine genotype, snake genotype, bovine genotype, etc. Nevertheless, this is all work in progress. To reinstate some stability to some of the major and clinically important genotypes, backed up by epidemiological data and experimental information, some were logically named as new species. Thus, the status quo of Upton and Current (1985) was abandoned and the following consensus was practically applied to cattle species. The abomasum is parasitized by C. andersoni and at least two named species are recognized to affect the intestine of cattle, the hostspecific C. bovis and the zoonotic C. pestis (Šlapeta, 2006). This approach logically aligns with the recognition of C. hominis which, along with C. pestis, were formerly classified as human and bovine genotypes of C. parvum.

Centenary of the Genus Cryptosporidium

47

Nevertheless this opinion is capable of reverting to the former conservative opinion, thus justifying the continuity of the names according to the ICZN, i.e. grouping C. hominis, C. parvum and C. pestis under the oldest available name, which is C. parvum, thereby reinstating the terms ‘human genotype’, ‘mouse genotype’ and ‘bovine genotype’, respectively. Underlying genetic differences have the potential to be used to distinguish between multiple taxa. While morphologically it is hard to find any differences, and host specificity provides only a partial picture, the use of DNA provides a virtual but very specific and repeatable label for each individual. If even a minor genetic difference is detected then it may reflect the niche adaptation, and so should be viewed as a practical tag for identification. Hence, those populations causing significant disease will stand out from the crowd and ultimately acquire a unique name. Indeed, the name C. hominis is now used for the dominant species transmitted from human to human, unequivocally typified by the draft genome of TU502 (Xu et al., 2004). Similarly, C. pestis is the name for the already known zoonotic species for which the draft genome of the Iowa strain is available (Abrahamsen et al., 2004). With the acceptance of both C. pestis and C. hominis we are clarifying the identity of the medically and veterinary important species. The synopsis for the three species is as follows: Cryptosporidium parvum Tyzzer, 1912 Syn. Cryptosporidium parvum ‘mouse genotype’ Xiao et al. (2002, 2004) Type host: Laboratory mouse, Mus musculus Type locality: Harvard Laboratory, Massachusetts, USA Cryptosporidium hominis Morgan-Ryan, Fall, Ward, Hijjawi, Sulaiman, Fayer, Thompson, Olson, Lal & Xiao, 2002 Syn. Cryptosporidium parvum ‘human genotype’ Xiao et al. (2002) Type host: Human, Homo sapiens Type locality: Perth, Western Australia, Australia Cryptosporidium pestis Šlapeta, 2006 Syn. Cryptosporidium parvum ‘bovine genotype’ Xiao et al. (2002, 2004) Type host: Domestic cattle, Bos taurus Type locality: Iowa, USA

Future Challenges Historically, two morphologically distinct populations of parasites of the gastrointestinal tract were originally described by Ernest E. Tyzzer from mice (i.e. Cryptosporidium muris and C. parvum). Nowadays, DNA sequences are increasingly seen as primary information sources for species identification in many organism groups, including Cryptosporidium. Such approaches stand on the implicit assumption that the reference databases used for comparison are sufficiently complete and feature-rich, with annotated entries. However, the uncertain taxonomic reliability and lack of annotations in public DNA repositories form a major obstacle to sequence-based species identification. The current taxonomic

48

J. Šlapeta

expansion of the genus Cryptosporidium is important relative to the dominant host, pathogenicity and genetic diversity. Unfortunately only few taxonomically available and named species have so far been molecularly characterized which, however, does not invalidate them from scientific usage. On the other hand, data from molecular epidemiological studies constantly reveal new genetic variants (= genotypes), but the lack of sufficient annotation and additional biological characteristics prohibits their elevation to species and ultimately taxonomic recognition. Nevertheless, descriptions of new genotypes remain an essential part of our understanding of the host specificity, the diversity and the epidemiology of the genus Cryptosporidium. In addition to the poor annotation of sequences in public DNA repositories, a huge gap exists between the number of described names and the number of identified genotypes. The closure of this gap represents a prime challenge for the decades to come.

Acknowledgements I would like to thank the organizers of the Second International Giardia and Cryptosporidium Conference (II IGCC) for their invitation to present the paper which forms the basis for this chapter. Financial support for the preparation of this article and participating in the II IGCC was partly provided by the Faculty of Veterinary Science, University of Sydney. I also thank Professor John Ellis (University of Technology Sydney) for editorial suggestions. I apologize to those authors whose work could not be cited owing to space limitations.

References 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. Awad-el-Kariem, F.M., Robinson, H.A., Dyson, D.A., Evans, D., Wright, S., Fox, M.T. and McDonald, V. (1995) Differentiation between human and animal strains of Cryptosporidium parvum using isoenzyme typing. Parasitology 110, 129–132. 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. Zeitschrift für Parasitenkunde 44, 289–298. Bednarska, M., Bajer, A., Kulis, K. and Sinski, E. (2003) Biological characterisation of Cryptosporidium parvum isolates of wildlife rodents in Poland. Annals of Agricultural and Environmental Medicine 10, 163–169. Bird, R.G. and Smith, M.D. (1980) Cryptosporidiosis in man: parasite life cycle and fine structural pathology. Journal of Pathology 132, 217–233. Casemore, D.P., Sands, R.L. and Curry, A. (1985) Cryptosporidium species a 'new' human pathogen. Journal of Clinical Pathology 38, 1321–1336. Current, W.L. and Reese, N.C. (1986) A comparison of endogenous development of three isolates of Cryptosporidium in suckling mice. Journal of Protozoology 33, 98–108.

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Duszynski, D.W. (1999) Revisiting the code: clarifying name-bearing types for photomicrographs of protozoa. Journal of Parasitology 85, 769–770. Fall, A., Thompson, R.C.A., Hobbs, R.P. and Morgan-Ryan, U.M. (2003) Morphology is not a reliable tool for delineating species within Cryptosporidium. Journal of Parasitology 89, 399–402. Fayer, R. and Ungar, B.L. (1986) Cryptosporidium and cryptosporidiosis. Microbiological Reviews 50, 458–483. Fayer, R., Santín, M. and Xiao, L. (2005) Cryptosporidium bovis n. sp. (Apicomplexa: Cryptosporidiidae) in cattle (Bos taurus). Journal of Parasitology 91, 624–629. Harris, D. (2003) Can you bank on GenBank? Trends in Ecology and Evolution 18, 317–319. International Commission on Zoological Nomenclature (1999) International Code of Zoological Nomenclature (ICZN), London, 306 pp. Available at: http://www.iczn.org/ iczn/index.jsp Iseki, M. (1979) Cryptosporidium felis sp. n. (Protozoa: Eimeriorina) from the domestic cat. Japanese Journal of Parasitology 28, 285–307. 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. Applied and Environmental Microbiology 61, 3849–3855. Khramtsov, N.V., Tilley, M., Blunt, D.S., Montelone, B.A. and Upton, S.J. (1995) Cloning and analysis of a Cryptosporidium parvum gene encoding a protein with homology to cytoplasmic form Hsp70. Journal of Eukaryotic Microbiology 42, 416–422. Kim, K., Gooze, L., Petersen, C., Gut, J. and Nelson, R.G. (1992) Isolation, sequence and molecular karyotype analysis of the actin gene of Cryptosporidium parvum. Molecular and Biochemical Parasitology 50, 105–113. Klesius, P.H., Haynes, T.B. and Malo, L.K. (1986) Infectivity of Cryptosporidium sp. isolated from wild mice for calves and mice. Journal of the American Veterinary Medical Association 189, 192–193. Korich, D.G., Marshall, M.M., Smith, H.V., O’Grady, J., Bukhari, Z., Fricker, C.R., Rosen, J.P. and Clancy, J.L. (2000) Inter-laboratory comparison of the CD-1 neonatal mouse logistic dose-response model for Cryptosporidium parvum oocysts. Journal of Eukaryotic Microbiology 47, 294–298. LeBlancq, S.M., Khramtsov, N.V., Zamani, F., Upton, S.J. and Wu, T.U. (1997) Ribosomal RNA gene organization in Cryptosporidium parvum. Molecular and Biochemical Parasitology 90, 463–478. Levine, N.D. (1984) Taxonomy and review of the coccidian genus Cryptosporidium (Protozoa, Apicomplexa). Journal of Protozoology 31, 94–98. Morgan, U.M., Constantine, C.C., O’Donoghue, P., Meloni, B.P., O’Brien, P.A. and Thompson, R.C. (1995) Molecular characterization of Cryptosporidium isolates from humans and other animals using random amplified polymorphic DNA analysis. American Journal of Tropical Medicine and Hygiene 52, 559–564. 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. (1999) The Cryptosporidium “mouse” genotype is conserved across geographic areas. Journal of Clinical Microbiology 37, 1302–1305. Morrison, D.A. (2006) Phylogenetic analyses of parasites in the new millennium. Advances in Parasitology 63, 1–124. Nilsson, R., Ryberg, M., Kristiansson, E., Abarenkov, K., Larsson, K. and Koljalg, U. (2006) Taxonomic reliability of DNA sequences in public sequence databases: a fungal perspective. PLoS ONE 1, e59.

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J. Šlapeta Ryan, U.M., Xiao, L., Read, C., Zhou, L., Lal, A.A. and Pavlasek, I. (2003) Identification of novel Cryptosporidium genotypes from the Czech Republic. Applied and Environmental Microbiology 69, 4302–4307. 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. Veterinary Parasitology 122, 103–117. Šlapeta, J. (2006) Cryptosporidium species found in cattle: a proposal for a new species. Trends in Parasitology 22, 469–474. Šlapeta, J. (2007) Response to Xiao et al.: Further debate on the description of Cryptosporidium pestis. Trends in Parasitology 23, 42–43. Spano, F., Putignani, L., McLauchlin, J., Casemore, D.P. and Crisanti, A. (1997) PCRRFLP 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 Microbiology Letters 150, 209–217. Templeton, T.J., Lancto, C.A., Vigdorovich, V., Liu, C., London, N.R., Hadsall, K.Z. and Abrahamsen, M.S. (2004) The Cryptosporidium oocyst wall protein is a member of a multigene family and has a homolog in Toxoplasma. Infection and Immunity 72, 980–987. Tilley, M., Upton, S.J. and Freed, P.S. (1990) A comparative study on the biology of Cryptosporidium serpentis and Cryptosporidium parvum (Apicomplexa: Cryptosporidiidae). Journal of Zoo and Wildlife Medicine 21, 463–467. Tyzzer, E.E. (1907) A sporozoan found in the peptic glands of the common mouse. Proceedings of the Society for Experimental Biology and Medicine 5, 12–13. Tyzzer, E.E. (1910) An extracellular coccidium, Cryptosporidium muris (gen. et sp. nov.) of the gastric glands of the common mouse. Journal of Medical Research 23, 487–509. Tyzzer, E.E. (1912) Cryptosporidium parvum (sp. nov.), a coccidium found in the small intestine of the common mouse. Archiv fur Protistenkunde 26, 394–412. Tzipori, S., Angus, K.W., Campbell, I. and Gray, E.W. (1980) Cryptosporidium: evidence for a single-species genus. Infection and Immunity 30, 884–886. Upton, S.J. and Current, W.L. (1985) The species of Cryptosporidium (Apicomplexa: Cryptosporidiidae) infecting mammals. Journal of Parasitology 71, 625–629. 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. Journal of Protozoology 18, 243–247. Xiao, L., Sulaiman, I.M., Ryan, U.M., Zhou, L., Atwill, E.R., Tischler, M.L., Zhang, X., Fayer, R., Lal, A.A. (2002) Host adaptation and host–parasite co-evolution in Cryptosporidium: implications for taxonomy and public health. International Journal for Parasitology 32, 1773–1785. Xiao, L., Fayer, R., Ryan, U. and Upton, S.J. (2004) Cryptosporidium taxonomy: recent advances and implications for public health. Clinical Microbiological Reviews 17, 72–97. Xiao, L., Fayer, R., Ryan, U. and Upton, S.J. (2007) Response to the newly proposed species Cryptosporidium pestis. Trends in Parasitology 23, 41–42. 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.

5

Molecular Epidemiology of Human Cryptosporidiosis in Developing Countries L. XIAO Centers for Disease Control and Prevention, Atlanta, GA, USA

Abstract Genotyping and subtyping tools have been used to characterize the transmission of human cryptosporidiosis in developing countries. Thus far, five Cryptosporidium spp. – C. hominis, C. parvum, C. meleagridis, C. canis and C. felis – are responsible for most Cryptosporidium infections in both immunocompetent and immunocompromised individuals. In most areas, C. hominis is responsible for over 70% of human cryptosporidiosis cases, with C. parvum accounting for 10–20% of infections. Differences have been observed among endemic areas in the proportion of infections due to each species, and in some areas C. meleagridis is also endemic. Results of subtyping suggest that there is high genetic heterogeneity in C. hominis in developing countries, and in these areas, human infections with C. parvum and other species are mostly the result of anthroponotic rather than zoonotic transmission. There is geographical segregation in C. hominis or C. parvum subtypes. Mixed and sequential infections with different Cryptosporidium species/genotypes and subtypes are common. Differences in oocyst shedding and clinical presentation have been observed among Cryptosporidium species and C. hominis subtypes. These findings reveal the diversity of cryptosporidiosis transmission in endemic areas and highlight the need for more extensive studies of cryptosporidiosis epidemiology in diverse areas with a wide spectrum of socioeconomic and environmental conditions.

Introduction Cryptosporidiosis is prevalent in developing countries, perhaps due to the high intensity of environmental contamination and poor hygiene conditions. Children in these highly endemic areas develop cryptosporidiosis very early in life, with peak infections usually occurring before 2 years of age. There is usually a strong association between the occurrence of cryptosporidiosis and the rainy season in tropical areas, or the cooler months in dry areas. These countries often have less intensive animal husbandry. Thus, the transmission of cryptosporidiosis in developing countries is probably different from that in the industrialized nations. Even though most molecular epidemiological studies of cryptosporidiosis have © CAB International 2009. Giardia and Cryptosporidium: From Molecules to Disease (eds G. Ortega-Pierres et al.)

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been conducted in the industrialized nations, an increasing number of studies are being done in developing countries. This has significantly improved our understanding of cryptosporidiosis transmission in these areas. This includes a better knowledge of parasite diversity, the roles of various transmission routes in cryptosporidiosis epidemiology, and the significance of parasite genetics in pathogenesis and clinical presentations. These developments have enabled health officials to better educate the public about the risk factors involved in the acquisition of cryptosporidiosis in vulnerable populations.

Molecular Epidemiological Tools Genotyping tools Many genotyping tools have been used to differentiate between Cryptosporidium species in humans. The PCR primers of earlier tools were mostly based on antigenic, structural, housekeeping genes and unknown genomic fragments of C. parvum, and included various formats of detection and differentiation (Xiao and Ryan, 2004; Cacciò, 2005). With few exceptions, most of these techniques can efficiently differentiate C. parvum, C. hominis and perhaps C. meleagridis, but are unlikely to amplify some of the more distant species (such as C. canis, C. felis, C. muris and C. andersoni). Therefore, these genotyping tools are mostly replaced by the genus-specific PCR-RFLP techniques based on the SSU rRNA gene, which have higher sensitivity and allow broad species detection.

Subtyping tools Subtyping tools are increasingly used in epidemiological studies of C. parvum and C. hominis. Several types of genetic targets are used in the development of subtyping tools, including microsatellites and minisatellites, double-stranded (ds) RNA elements, and ITS-2. In developing countries, the most widely used subtyping tool is DNA sequencing of the 60 kDa glycoprotein (GP60) gene. The GP60 gene 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 non-repeat 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 (Fig. 5.1). 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). It should be kept in mind that GP60 and other subtyping tools (including the MST and MLST tools described below) do not amplify the DNA of C. felis, C. canis and other species distant from C. parvum and C. hominis.

Molecular Epidemiology of Human Cryptosporidiosis in Developing Countries 100

IIi-Akiyoshi-154

100

0.1 substitutions/site

53

Id-AF164497 I-Akiyoshi-90

100

70

II-Kenya-9589R IIe-7870

82

II-Alpaca-13987 IIj-NI-8934 IIf-7490 I-Peru9911

100 98

Ia-AF164502 Ig-NI3986

99

IId-4833 IIb-4736

99 100

IIg-Akiyoshi-66

100

FelisAY700394.1| 99

IIa-4804: Zoonotic C. parvum Ib-4746

100

Ie-5632 67

If-4755

55 IIh-Akiyoshi-112

52

II-kenya-9408R

67

IIc-4742: Anthroponotic C. parvum III-DQ067570

100 98

III-4500 III-295

Fig. 5.1. Phylogenetic relationships between the known subtype families of C. hominis (Ia, Ib, Id, Ie, If, etc.) and C. parvum (IIa, IIc, IId, IIe, etc.) based on a neighbour-joining analysis of the GP60 gene sequences. Sequences starting with III were derived from C. meleagridis. Two major anthroponotic and zoonotic C. parvum subtype families (shown in boxes) are identified.

MLT and MLST The recent genome sequencing of C. parvum and C. hominis has allowed the identification of microsatellite and minisatellite sequences in C. parvum and C. hominis genomes and other targets that are highly polymorphic between C. parvum and C. hominis. They are frequently used in multilocus analysis to increase the typing resolution. Two types of techniques are used in typing. In multilocus typing (MLT), variation in microsatellites and minisatellites is assessed on the basis of length variations, using polyacrylamide gel electrophoresis or the GeneScan technology (Ngouanesavanh et al., 2006). This allows the use of

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many targets in the MLT techniques economically. The second kind of typing technique, multilocus sequence typing (MLST), relies on the detection of genetic heterogeneity by DNA sequencing of the amplified PCR products (Gatei et al., 2006a, 2007). Compared to MST, MLST allows the detection of nucleotide substitution and the inclusion of markers with single nucleotide polymorphisms.

Proper use of molecular epidemiological tools Caution should be exercised in interpreting typing results, as accurate diagnosis of species and subtypes has important implications for understanding infection sources. In two studies in India, Cryptosporidium mouse genotype was identified in one child and one HIV-positive adult, based on results of RFLP analysis of the SSU rRNA PCR products (Muthusamy et al., 2006; Ajjampur et al., 2007). The true identity of the parasite was probably C. meleagridis, which has the same RFLP pattern as the mouse genotype in the genotyping technique used. The detection of C. andersoni in a Malawian child was also largely based on RFLP analysis (Morse et al., 2007), which needs to be confirmed by DNA sequencing, as DraI and AseI RFLP was mainly used in species differentiation and C. muris may also share the same pattern. The use of RFLP instead of DNA sequencing in differentiating GP60 subtypes should be avoided, as some subtype families share high sequence homology in one region but not in other regions of the gene, and many new subtype families have been found since the initial development of the RFLP differentiation tool. The sequence mosaic in the gene is also responsible for the polyphyletic nature of both C. parvum and C. hominis in the gene (Fig. 5.1). Thus, the identification of IIa and If subtype families in humans in southern India (Muthusamy et al., 2006) needs to be confirmed by the results of DNA sequencing. The subtype family name Ic is still occasionally used by some researchers to describe C. parvum (Muthusamy et al., 2006; Ajjampur et al., 2007), even though this was the result of an initial misidentification of species at the SSU rRNA locus (Strong et al., 2000), and the subtype family was renamed as IIc some time ago (Alves et al., 2003). IIc should be used to avoid unnecessary confusion. In one study (Ajjampur et al., 2007), the so-called Cryptosporidium mouse genotype GP60 sequence was probably from C. meleagridis, and the C. felis GP60 sequences were probably not from C. felis.

Cryptosporidium Species and Genotypes in Humans In a similar way as for people living in the industrialized nations, five Cryptosporidium spp. are responsible for most human Cryptosporidium infections in developing countries; including C. hominis, C. parvum, C. meleagridis, C. canis and C. felis. They were initially found in otherwise healthy children in Peru in a longitudinal cohort study using a SSU rRNA-based genotyping tool (Xiao et al., 2001), but have also recently been found in diarrhoeic children in Kenya (Gatei et al., 2006b). One large-scale study of Ugandan children reported only

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Table 5.1. Distribution of common human-pathogenic Cryptosporidium species in children in developing countries. No. of isolate C. hominis C. parvum C. meleagridis C. felis C. canis Reference

Location China India India Kenya Malawi Malawi Uganda Iran Guatemala Lima, Peru Chile

5 50 58

5 47 47

0 0 7

175 37 43 444

153 35 25 326

15 2 10 85

7

3

4

15 85 4

14 67 2

1 8 2

1?

1 3

1

2

3

1

2

2 5

7

Peng et al. (2001) Gatei et al. (2007) Ajjampur et al. (2007) Gatei et al. (2006b) Peng et al. (2003) Morse et al. (2007) Tumwine et al. (2003) Meamar et al. (2007) Xiao et al. (2004) Xiao et al. (2001) Neira-Otero et al. (2005)

C. hominis, C. parvum and C. meleagridis, but the genotyping tool used is not capable of detecting C. canis, C. felis and other species genetically distant from C. parvum and C. hominis (Tumwine et al., 2003). Other studies examined only small numbers of specimens, which was probably responsible for the low species diversity detected (Table 5.1). These five Cryptosporidium spp. have also been found in immunocompromised people in developing countries (Table 5.2). Studies conducted in Thailand, India and Peru showed the presence of these five species in HIV-positive adults (Gatei et al., 2002b; Cama et al., 2003). Small-scale studies in other areas have also revealed the presence of some of the five species in HIV-positive adults or children (Table 5.2). In Lima, Peru, there is no significant difference in the distribution of the five species between children and HIV-positive adults (Xiao et al., 2001; Cama et al., 2003). It is likely that other Cryptosporidium species can infect humans under certain circumstances. Cryptosporidium muris has been found in human cases in Indonesia (Katsumata et al., 1998), Kenya (Gatei et al., 2002a, 2006b), Peru (Palmer et al., 2003) and India (Muthusamy et al., 2006). Other Cryptosporidium species found in humans in developing countries include C. suis in one HIVpositive adult and Cryptosporidium cervine genotype in a child, both in Lima, Peru (Xiao et al., 2002; V. Cama et al., unpublished). New Cryptosporidium genotypes will probably be found in humans in future, even though these parasites account for a very small proportion of Cryptosporidium infections in humans.

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Table 5.2.

Distribution of common human-pathogenic Cryptosporidium species in HIV-positive people in developing countries.

Location

Age

Vellore, India Bangkok, Thailand

Adults 4 children, 25 adults Adults Adults Adults Children Children Children Adults Adults Mostly adults Adults Adults

Bangkok, Thailand Taiwan Kenya Malawi Ugandaa South Africa Iran Venezuela Colombia Haiti Lima, Peru a Including

No. of isolate C. hominis C. parvum C. meleagridis C. felis C. canis Reference 48 29

31 24

9

1 + 1 (?) 3

5 1

34 4 24 6 76 21 8 10 6 49 302

17 2 14 6 56 16 1 8 3 31 204

5

7 1 1

3 1

8 14 5 7 1 2 16 34

Muthusamy et al. (2006) Tiangtip and Jongwutiwes (2002) 2

3

1 1? 38

1 1? 10

12

Gatei et al. (2002b) Hung et al. (2007) Gatei et al. (2003) Peng et al. (2003) 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)

nine HIV-negative children.

L. Xiao

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Cryptosporidium hominis Infection In developing countries, more than 70% of Cryptosporidium infections in children are caused by C. hominis in most of the areas studied (Table 5.1). In these areas, children acquire cryptosporidiosis very early in life, with peak incidence of infections occurring between 1 and 2 years of age. Under such conditions, the transmission of cryptosporidiosis is probably mostly through a direct person-to-person route rather than an indirect waterborne or foodborne route. This is probably responsible for the high proportion of infections attributable to C. hominis. This predominance of C. hominis in humans in developing countries applies also to HIV-infected adults (Table 5.2). A diarrhoea outbreak caused by C. hominis also occurred in a daycare centre in São Paulo, Brazil (Goncalves et al., 2006). Molecular characterizations of C. hominis isolates have revealed the complexity of cryptosporidiosis epidemiology. Despite the seemingly lower diversity of Cryptosporidium in humans at the species/genotype level in developing countries, the results of GP60 subtyping have revealed the complexity of Cryptosporidium transmission in endemic areas. This is evident from the existence of many C. hominis subtype families in each endemic area. Thus, 3–4 C. hominis subtype families were seen in humans in India, Peru, Kenya, Malawi and South Africa (Leav et al., 2002; Peng et al., 2003; Xiao et al., 2004; Ajjampur et al., 2007; Cama et al., 2007; Gatei et al., 2007). In these areas, the complexity of transmission is frequently reflected by the existence of many subtypes within C. hominis subtype families Ia and Id in one endemic area (Leav et al., 2002; Peng et al., 2003; Cama et al., 2007; Gatei et al., 2007). The high C. hominis heterogeneity in developing countries is probably an indicator of intensive and stable cryptosporidiosis transmission in the area. Four common C. hominis subtype families – Ia, Ib, Id and Ie – have been found in humans in many developing countries (Xiao et al., 2004) (Fig. 5.1). Nevertheless, there are geographical differences in their distribution. For example, Ia, Ib and Id are common in Malawi, South Africa, India and Peru (Leav et al., 2002; Peng et al., 2003; Ajjampur et al., 2007; Cama et al., 2007; Gatei et al., 2007). 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). In contrast, the common subtype family Ie was not seen in cryptosporidiosis in South African children (Leav et al., 2002). Two space–time clusters of Ia infections were identified in children in southern India (Ajjampur et al., 2007), but it is unclear whether each cluster was caused by one Ia subtype, as many subtypes of the Ia subtype family are usually present in an area (see above). 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 C. hominis subtype family Ib: IbA9G3 and IbA10G2. The former is commonly seen in Malawi, Kenya and India, whereas the latter is commonly seen in South Africa, Botswana, Jamaica and Peru (Leav et al., 2002; Cama et al., 2007; Gatei et al., 2006a, 2007; L. Xiao et al., unpublished). This geographical segregation of C. hominis subtypes becomes much more obvious when specimens from different countries are genetically compared using a MLST tool (Gatei et al., 2006a).

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Cryptosporidium parvum Infection Most developing countries have very low frequency of C. parvum infections, although the burden attributable to this species seems to vary by area. In these countries, C. parvum transmission in humans is frequently anthroponotic. This is supported by results of GP60 subtyping in several regions. Two major GP60 subtype families, the zoonotic IIa and the anthroponotic IIc, are responsible for most C. parvum infections in humans (Fig. 5.1). IIa subtypes are rarely seen in humans in developing countries. Instead, the subtype family IIc is responsible for most human C. parvum infections in these areas (Leav et al., 2002; Peng et al., 2003; Xiao et al., 2004; Akiyoshi et al., 2006; Ajjampur et al., 2007). In some places, such as Lima, Peru and southern India, 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., 2003; Xiao et al., 2004; Ajjampur et al., 2007; Cama et al., 2007). In Uganda, even though IIc subtypes are the dominant C. parvum in children, several new subtype families have been reported (Akiyoshi et al., 2006). All these are likely indicators of the reduced role of zoonotic transmission of C. parvum in these areas. Nevertheless, a recent study of asymptomatic cryptosporidiosis in Zambian dairy farm workers and their household members has identified C. parvum as being more common than C. hominis (Siwila et al., 2007). Zoonotic infection was also suspected to be one cause of the seemingly wider species diversity in Malawian children living in rural areas (Morse et al., 2007). No subtyping of C. parvum was done in these studies to identify possible infection sources. One major outbreak of diarrhoea largely attributed to cryptosporidiosis occurred in Botswana in early 2006 after heavy rain, causing thousands of infections and several hundreds of deaths in a number of districts. Both C. parvum and C. hominis were identified in infected people, with the former responsible for more cases. As expected, five C. hominis subtype families and six subtypes were identified in a small number of specimens analysed. However, the C. parvum specimens analysed had only IIcA5G3a or IIcA5G3b subtypes of the IIc subtype family (L. Xiao et al., unpublished). Because C. parvum was identified as the major species and some patients in the outbreak had enteropathogenic E. coli, cattle waste was initially suspected as the major source of contamination. The finding of only the C. parvum subtype IIc family and the presence of C. hominis clearly suggests that the major initial source of contamination was human sewage.

Infections with Other Species/Genotypes In most developing countries studied, C. parvum and C. hominis are responsible for more than 90% of human cases of cryptosporidiosis, with the remainder attributable to C. meleagridis, C. canis and C. felis. Some areas, however, have a high prevalence of these unusual species. In Lima (Peru) and Bangkok (Thailand), C. meleagridis is as prevalent in humans as C. parvum, being responsible for

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10–20% of human cryptosporidiosis cases (Xiao et al., 2001; Gatei et al., 2002b; Cama et al., 2003). Subtyping tools are largely unavailable for these rare species because most C. parvum- and C. hominis-based subtyping primers are not capable of PCR amplification in these species. GP60 and HSP70-based typing tools, however, are available to subtype C. meleagridis. Multiple C. meleagridis subtypes were seen in the small number of human specimens from several different areas (Glaberman et al., 2001). A recent MLST analysis in a small community in Lima, Peru, has shown the presence of multiple subtypes in children, AIDS patients, and birds, with no apparent host segregation of the subtypes found (L. Xiao et al., unpublished). Like C. parvum, most human infections with the usual species are probably the result of anthroponotic transmission, judged by the concurrent presence of C. hominis in some of the C. meleagridis-, C. canis- or C. felis-infected individuals (see below). Possible transmission of C. canis between two siblings and a dog occurred in a household in a slum in Lima, Peru, but the direction of the transmission was not clear (Xiao et al., 2007).

Mixed Infections and Sequential Infections No matter which PCR tool is used in genotyping Cryptosporidium, all broadly specific tools have the problem that they detect 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 for them 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 probably under-detected by these tools (Reed et al., 2002). A low prevalence of mixed infections of multiple Cryptosporidium genotypes has been reported in children in Uganda (19/444), Malawi (1/43) and India (2/50) (Tumwine et al., 2003; Gatei et al., 2007; Morse et al., 2007). Using genotype-specific primers in combination with other genus-specific primers, Cama et al. (2006) identified concurrent C. hominis, C. parvum or C. meleagridis infections in seven out of 21 C. canis- or C. felis-infected HIV-positive people. Seven out of 55 C. meleagridis-infected children and HIV-positive people in Lima, Peru, had concurrent C. hominis infection in a MLST analysis (L. Xiao et al., unpublished). Thus, the accurate identification of infections with mixed genotypes has important implications for understanding the transmission of so-called zoonotic species and genotypes in humans. In developing countries, children frequently have multiple episode of cryptosporidiosis, although the likelihood of clinical illness decreases with increased infection episodes (Bern et al., 2002). Molecular analysis of longitudinal samples from Peruvian children with multiple cryptosporidiosis episodes indicated that immunity against both homologous and heterologous Cryptosporidium species/ genotypes was short-lived, with time intervals between infections of about 1 year. As might be expected, sequential infections with heterologous Cryptosporidium species were more common than sequential infections with homologous

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L. Xiao

Cryptosporidium species (Xiao et al., 2001, Cama et al., 2008). Even though some children had the same genotype (C. hominis) in sequential infections, results of subtyping suggested that these were in fact due to heterogeneous subtypes of the parasite (Cama et al., 2008).

Cryptosporidium Species/Genotypes, Subtypes, and Virulence The clinical and epidemiological significance of various Cryptosporidium species and subtypes in humans is not yet clear. Results of recent genotyping studies nevertheless support the theory that C. hominis and C. parvum behave differently in humans. Studies in slums in Peru and Brazil have shown that children infected with C. hominis have higher oocyst shedding intensity and longer duration than those infected with C. parvum and other genotypes (Xiao et al., 2001; Bushen et al., 2007). Children infected with C. hominis had a significantly greater severity of diarrhoea than those infected with other species in southern India (Ajjampur et al., 2007). In AIDS patients in Lima, Peru, only infections with C. canis, C. felis or subtype family Id of C. hominis were significantly associated with diarrhoea, and infections with C. parvum were significantly associated with chronic diarrhoea and vomiting. Infections with C. hominis Ib subtype family were also marginally associated with diarrhoea and vomiting. In contrast, infections with C. meleagridis and Ia and Ie subtype families of C. hominis were usually asymptomatic (Cama et al., 2007). These results demonstrate that different Cryptosporidium genotypes and subtype families are linked to different clinical manifestations. Because of the differences in pathogenicity, C. hominis and C. parvum seemingly have different nutritional effects on infected children. In Brazil, heightfor-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).

Conclusions Molecular epidemiological studies of cryptosporidiosis in developing countries are still in their infancy, but significant progress has already been made towards a better understanding of the transmission of the infection. We are beginning to use second-generation molecular tools to answer epidemiological questions that are difficult to address by traditional methods, such as maintenance of immunity and cross-protection, transmission dynamics in different settings, temporal and geographical variations in Cryptosporidium transmission, and the role of parasite factors in the variability in transmission and clinical spectrum of cryptosporidiosis. More importantly, we are starting to see better collaboration between epidemiologists, clinicians, molecular biologists and parasitologists in well-designed

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epidemiological studies. Such an integrated approach will undoubtedly lead to better utilization of the available molecular diagnostic tools and a better understanding of the epidemiology of cryptosporidiosis.

Acknowledgements The findings and conclusions reported in this chapter are those of the author and do not necessarily represent the views of the Centers for Disease Control and Prevention.

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Molecular Epidemiology and Typing of Non-human Isolates of Cryptosporidium U.M. RYAN1 AND L. XIAO2 1Murdoch

University, WA, Australia; 2Center for Disease Control and Prevention, Atlanta, GA, USA

Abstract Cryptosporidium has been reported in a wide variety of hosts, with C. parvum and C. hominis being responsible for most human infections. Until recently, it has been assumed that farm animals and wild animals are important zoonotic reservoirs for human cryptosporidiosis. However, recent molecular analysis has revealed a wide range of Cryptosporidium species and genotypes infecting both domestic and wild animals, and the epidemiology of cryptosporidiosis is clearly more complicated than was previously thought.

Introduction Cryptosporidium has been reported in a wide variety of vertebrate hosts (Fayer et al., 2000) and, at present, 16 species of Cryptosporidium are regarded as valid on the basis of differences in oocyst morphology, site of infection, vertebrate class specificity and genetic differences: C. muris in rodents; C. andersoni and C. bovis in cattle and sheep; C. suis in pigs; C. parvum in cattle, humans and other mammals; C. meleagridis in birds and humans; C. hominis in humans; C. baileyi and C. galli in birds; C. serpentis and C. saurophilum in snakes and lizards; C. molnari and C. scophthalmi in fish; C. wrairi from guinea pigs; C. felis in cats; and C. canis in dogs (Fayer et al., 2000, 2001; Alvarez-Pellitero and Sitjà-Bobadilla, 2002; Morgan-Ryan et al., 2002; Ryan et al., 2003a, 2004a; Alvarez-Pellitero et al., 2004). Cryptosporidium parvum and C. hominis are responsible for most human infections (Morgan-Ryan et al., 2002), and it has been assumed that the majority of Cryptosporidium infections in farmed animals that had oocysts in the size range of 4–6 mm were due to C. parvum (cattle genotype) and that farm animals and wild animals represented an important zoonotic reservoir for human cryptosporidiosis. However, recent molecular analysis has revealed a wide range of Cryptosporidium species and genotypes infecting both domestic and wild animals, © CAB International 2009. Giardia and Cryptosporidium: From Molecules to Disease (eds G. Ortega-Pierres et al.)

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and the epidemiology of cryptosporidiosis is clearly more complicated than was previously thought.

Cryptosporidium Species and Genotypes in Sheep Cryptosporidium has been reported in sheep worldwide; however, most studies on Cryptosporidium in sheep have been based on microscopy, with the reported prevalence ranging from 2.6% to 82% (see Ryan et al., 2005). Recent molecular characterization of Cryptosporidium in sheep suggests that the majority of sheep are not infected with zoonotic genotypes. In Australia, a survey of 1647 sheep faecal samples reported a prevalence of 26.2%. The most common Cryptosporidium genotypes identified were the cervid genotype (55%) and C. bovis (23%). Low levels of C. andersoni, C. suis, marsupial genotype, pig II genotype, C. hominis and a novel previously unidentified genotype were also detected (Ryan et al., 2005). Cryptosporidium parvum was not detected in any of the samples. More recently, in the USA, sequence analysis of 57 specimens corresponding to 8 ewes and 24 lambs that were positive for Cryptosporidium identified the cervid genotype (84%), C. bovis-like genotype (~12%) and C. parvum (3.5%) (Santín et al., 2007). In the UK, C. parvum and a novel genotype were reported in sheep (Chalmers et al., 2002); however, RFLP analysis of the COWP locus was used (Chalmers et al., 2002) and recent studies have shown that this analysis can erroneously identify isolates as C. parvum due to the extent of conservation at the COWP locus (Hamnes et al., 2007).

Cryptosporidium Species and Genotypes in Pigs Cryptosporidium was first reported in pigs in the USA in 1977 (Kennedy et al., 1977). Subsequent studies have been undertaken worldwide, including Australia, Japan, Germany, Spain and Ireland, with prevalences ranging from 1.4% to 100% in pigs aged 1 week to adulthood (Quílez et al., 1996; Izumiyama et al., 2001; Wieler et al., 2001; Maddox-Hyttel et al., 2006; Xiao et al., 2006). The prevalence in piglets under 2 months of age has been reported to range between 0% and 59.2%. The age at which Cryptosporidium is most prevalent is in pigs between 6 and 12 weeks old (see Hamnes et al., 2007). The majority of nursing piglets and weanlings that have been positive for Cryptosporidium have been shown to be infected with two Cryptosporidium spp. – C. suis and pig genotype II (Guselle et al., 2003; Ryan et al., 2003a, 2004a; Vitovec et al., 2006; Xiao et al., 2006; Hamnes et al., 2007; Langkjær et al., 2007) (see Table 6.1). Cryptosporidium suis is adapted to porcine hosts, but poorly infective for cattle (Enemark et al., 2003) and not infective for nude mice and neonatal BALB/c mice (Morgan et al., 1999a; Ryan et al., 2003a; Vitovec et al., 2006). Cryptosporidium suis has been identified in humans (Xiao et al., 2002a) but is not a common human pathogen. Little is known about the pathogenicity or zoonotic potential of pig genotype II but it has not been reported in humans to date and is unlikely to be a major threat to public health. Phylogenetic analysis

Molecular Epidemiology and Typing of Non-human Isolates Table 6.1.

67

Cryptosporidium genotypes infecting pigs.

Country Denmark Weaners Piglets Norway Australia

No. genotyped

C. suis

Pig genotype II

170 13 9 12

24% 71% ~66% 60%

76% 29% ~33%

Other species

Reference Langkjær et al. (2007)

C. parvum (40%)

Australia Weaners Piglets Australia Weaners Piglets Ireland

14 14

50% 50%

50% 50%

27 18 25

24% 100% 57%

76% 0% 39%

Czech Republic

13

100%

Hamnes et al. (2007) Morgan et al. (1999a) Ryan et al. (2003a)

Johnson et al. (2008)

C. muris (3.5%)

Xiao et al. (2006) Vitovec et al. (2006)

at multiple loci has confirmed the species status of C. suis and provides strong evidence that pig genotype II is also a valid species (see Ryan et al., 2003a; Xiao et al., 2004a). In Denmark, sequence analysis of 183 Cryptosporidium-positive pig isolates from sows, weaners and piglets from various herds identified C. suis in 24% of weaners and 71% of piglets, and pig genotype II in 76% of weaners and 29% of piglets (Langkjær et al., 2007). Higher oocyst concentrations were observed in samples genotyped as C. suis than in samples genotyped as pig genotype II. There also appeared to be an age-related change in the species/genotypes infecting pigs, as pig genotype II was more prevalent in weaners whereas the majority of piglets were infected with C. suis (Langkjær et al., 2007). This is in contrast with a previous Australian study in which 28 Cryptosporidium isolates from pigs were genotyped, revealing equal numbers of C. suis and pig genotype II (Ryan et al., 2003a). No correlation between genotype and host age was found, but some pigs infected with pig genotype II seemed to have a high excretion of oocysts (Ryan et al., 2003a). However, a more recent study in Australia reported that pig genotype II was more prevalent in weaners (71%) than C. suis (17%) (Johnson et al., 2008). One of the first genotyping studies on pigs identified C. suis but also C. parvum (cattle genotype) by sequence analysis of the 18S rRNA gene in outdoor pigs suffering from diarrhoea in Western Australia (Morgan et al., 1999a). Interestingly, C. parvum has not been detected in pig isolates since then and it is likely that C. parvum is responsible for only occasional infections in pigs. In the Czech Republic, 0/135 sows, 193/3368 (5.7%) pre-weaned and 201/835 (24.1%) post-weaned piglets were positive for Cryptosporidium infection. Only C. suis was identified; however, only a few isolates were genotyped

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(Vitovec et al., 2006). The authors reported that the C. suis strains obtained were larger (6.2 × 5.5 mm) than the oocyst sizes given in the original C. suis description (4.6 × 4.2 mm) (Ryan et al., 2004a). However, studies have shown that there is substantial morphological variation in oocyst size within individual species and that morphology alone is not a reliable indicator for delimiting species (Fall et al., 2003). In Norway, 31 (31%) herds and 57 (8.3%) litters from 684 litters of suckling piglets from 100 indoor swine herds from all regions tested positive for Cryptosporidium. Molecular characterization of nine Cryptosporidium isolates demonstrated both C. suis and Cryptosporidium pig genotype II (Hamnes et al., 2007). A study in Ireland reported that the spreading of pig slurry onto pasture or crops may lead to increases in the occurrence of Cryptosporidium in water, as C. suis, Cryptosporidium pig genotype II, and C. muris were identified in 25 of 56 pig slurry samples from 33 Irish farms (Xiao et al., 2006). The finding of C. muris in pig slurry is unusual and could have originated from either infected pigs or rodents. Even though C. muris infection has never been reported in pigs, this species does have the widest host range of all Cryptosporidium species, and this range includes humans (Iseki et al., 1989).

Cryptosporidium Species and Genotypes in Cattle Since the late 1980s, cattle have been identified as being one of the main reservoir host for the zoonotic C. parvum; however, recent studies in the USA suggest that cattle are infected with at least four Cryptosporidium parasites: C. parvum, C. bovis, C. andersoni and the Cryptosporidium deer-like genotype (Santín et al., 2004; Fayer et al., 2006, 2007; Feng et al., 2007; Xiao et al., 2007). Recent studies in the USA indicated that the occurrence of these Cryptosporidium spp. in cattle is age-related (Santín et al., 2004), as the zoonotic C. parvum was responsible for about 85% of the Cryptosporidium infections in pre-weaned calves but only 1% of the Cryptosporidium infections in post-weaned calves and heifers (Santín et al., 2004). Post-weaned calves were mostly infected with C. bovis, C. andersoni and the deer-like genotype (Santín et al., 2004) (see Table 6.2). These findings clearly demonstrate that neonatal calves are an important source of zoonotic cryptosporidiosis in humans. Neonatal calves are also the age group of cattle mostly affected by cryptosporidiosis in terms of prevalence of infection and the associated morbidity and mortality (Fayer et al., 1997). Studies on heifers aged 12–24 months of age on dairy farms in Pennsylvania, Vermont, New York, Maryland, Virginia, North Carolina and Florida reported a much lower prevalence of C. parvum and reported that C. suis, C. parvum, the deer-like genotype, C. bovis and C. andersoni accounted for 1%, 6%, 15%, 35% and 43%, respectively (Fayer et al., 2006). A study on mature dairy cattle identified an even lower prevalence of Cryptosporidium and reported that C. parvum, C. bovis and C. andersoni were found infecting 0.4%, 1.7% and 3.7% of the 541 cows, respectively (Fayer et al., 2007). The overall lower prevalence of Cryptosporidium in these cows was very highly significant (P < 0.0001) compared

Cryptosporidium genotypes infecting cattle.

Country USA Pre-weaned Post-weaned USA Heifers USA Milking cows New York state Georgia, USA Pre-weaned Post-weaned Milking cows China India Denmark Cows Older calves Young calves Hungary Calves Portugal Calves Adults aC.

No. Genotyped

C. parvum

C. bovis

C. andersoni

Deer-like genotype

278

85% 1%

9% 55%

1% 13%

5% 31%

68

6%

35%

43%

15%

31 115

6.5% 61% (70/115)

29% 37% (42/115)

23 6 3 6 12

26% 0% 0%

3 61 90

100% 4% 82%

22

95%

63 7

100% 71.5%

Other species

Reference Santín et al. (2004)

Fayer et al. (2006) 1%a Fayer et al. (2007) 64.5% 3% (3/115)

Starkey et al. (2006) Feng et al. (2007)

8.4%

39% 66% 66% 83% 91.6% – 73% 14%

22% 16% 0%

33%b

16%

Feng et al. (2007) Feng et al. (2007) Langkjær et al. (2007)

– 14% 3%

– 4%c Plutzer and Karanis (2007) Mendonça et al. (2007)

5% 14.25%

Molecular Epidemiology and Typing of Non-human Isolates

Table 6.2.

14.25%d

suis; b1 mixed C. parvum + C. andersoni isolate; cC. suis-like genotype + <1% pig genotype II; dC. meleagridis. 69

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with younger cattle. The very low level of infection with zoonotic C. parvum, suggests that mature dairy cattle are a relatively low-risk source of infection for humans (Fayer et al., 2007). A study on 115 faecal samples positive for Cryptosporidium from dairy cattle in New York state, identified 70 of the 115 (61%) as C. parvum, 42 (37%) as C. bovis, and 3 (3%) as the deer-like genotype (Starkey et al., 2006). The authors also suggested that C. bovis may be more host-adapted and thus less pathogenic to dairy cattle than C. parvum (Starkey et al., 2006). The prevalence of Cryptosporidium species in cattle in other countries is less well characterized but recent studies have shown that C. bovis and the deer-like genotype are present in cattle studied in China and India (Feng et al., 2007) (see Table 6.2). In that study, the prevalence of Cryptosporidium species in cattle in Georgia was compared with that in China and India. Analysis revealed that the prevalence of C. bovis and the deer-like genotype was much higher than that of C. parvum, even in pre-weaned calves, in Georgia. This is in contrast to the studies by Fayer et al. (2006) and Santín et al. (2004), which reported that that C. bovis and the deer-like genotype were mostly found in post-weaned calves and older animals. This may have been due to the fact that the pre-weaned calves examined in the Feng study were mostly 1 month or older and only a small number of animals were sampled. Alternatively, cattle of all ages may be susceptible to infections with both species and there may not be an age-associated occurrence of the two parasites in cattle. It is also possible that calves may acquire infection with C. bovis or the deer-like genotype early in life, but the infection may be concealed by the overwhelming C. parvum infection. The pre-patent period for C. bovis is 10–12 days, which is much longer than for C. parvum (about 4 days) (Fayer et al., 2005). Further studies are required to elaborate on this. A study in Denmark also showed age structuring, with C. bovis causing 14% and 73% of Cryptosporidium infections in young and older calves, respectively (Langkjær et al., 2007). Cryptosporidium parvum and the deer-like genotype were responsible for 82% and 4% in young calves and 3% and 14% of Cryptosporidium infections in older calves (Langkjær et al., 2007). A novel C. suislike genotype which exhibited 99% and 98% nucleotide identity to C. suis at the 18S rDNA and HSP70 loci, respectively, was identified in three calves originating from two different herds (Langkjær et al., 2007). Pig genotype II, which until now has appeared to be pig-specific, was also identified in a young calf (Langkjær et al., 2007). Cryptosporidium andersoni, which was found previously in Danish cattle (Enemark et al., 2002), was not detected. This is probably explained by the fact that C. andersoni is primarily found in older animals and that only three isolates from cows could be successfully genotyped. In Portugal, a study of 467 dairy and beef cattle, of which 291 were calves and 176 were adults, identified 74/291 calves (25.4%) and 8/176 adults (4.5%) as positive for Cryptosporidium by microscopy (Mendonça et al., 2007). Molecular characterization of 63 isolates from calves and 7 isolates from adults, identified all of the isolates as C. parvum using sequence analysis of the HSP70 locus. However, sequence analysis of the 18S rRNA locus identified two of the C. parvum isolates as C. meleagridis and C. andersoni (Mendonça et al., 2007). This is the first report of C. meleagridis in cattle.

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In Hungary, 79 faecal samples from calves with diarrhoea were collected on 52 farms from different counties (Plutzer and Karanis, 2007). Immunofluorescence microscopy identified 39 samples as positive for Cryptosporidium and 18S rRNA RFLP analysis of 22 isolates identified C. parvum in 21 samples and the Cryptosporidium deer-like genotype in 1 sample (Plutzer and Karanis, 2007).

Cryptosporidium in Wild Animals Little is known of the prevalence and distribution of Cryptosporidium spp. infecting wild animals, but studies to date suggest that the majority of wild animals do not shed oocysts that are of public health significance. A recent study detected 36 Cryptosporidium positives from 471 faecal samples from foxes, raccoons, muskrats, otters and beavers living in wetlands adjacent to Chesapeake Bay, Maryland, USA. Genotyping analysis identified five types of Cryptosporidium, including the C. canis dog and fox genotypes, Cryptosporidium muskrat genotypes I and II, and Cryptosporidium skunk genotype (Zhou et al., 2004a). The Cryptosporidium ferret genotype has been identified in ferrets, river otters and American mink (Xiao et al., 2002b; Gaydos et al., 2007; Gomez-Couso et al., 2007). Another study examined faeces and duodenal scrapings from 22 coyotes killed in managed hunts in north-eastern Pennsylvania. Six coyotes (27%) were positive for Cryptosporidium spp.; one isolate shared 99.7% homology with C. muris, whereas five others (23%) shared 100% homology with C. canis coyote genotype (Trout et al., 2006). In marsupials, multiple species and genotypes have been identified, including marsupial genotype I and II, opossum genotype I (which is genetically very closely related to marsupial type I), opossum genotype II and C. muris (Morgan et al., 1997; Xiao et al., 1999a, 1999b, 2002b; Warren et al., 2003; Power et al., 2004, 2005). A recent study examined the occurrence of Cryptosporidium oocysts in faeces from a population of wild eastern grey kangaroos inhabiting a protected watershed in Sydney, Australia (Power et al., 2005). Over a 2-year period, Cryptosporidium oocysts were detected in 239 of the 3557 (6.7%) eastern grey kangaroo faecal samples tested by using a combined immunomagnetic separation and flow cytometric technique. The prevalence of Cryptosporidium in this host population was estimated to range from 0.32% to 28.5%, with peaks occurring during the autumn months. Partial 18S rRNA gene sequencing of 51 isolates representing 10 of the 11 faecal sampling periods identified marsupial genotypes I and II in 61% and 39% of samples, respectively (Power et al., 2005). As marsupial genotypes I and II have not been identified in humans to date, it is unlikely that marsupials in water catchment areas are a serious public health threat.

Cryptosporidium in Rodents There have been few studies conducted on the epidemiology of Cryptosporidium in wild rodents; however, it is important to establish whether they are a

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reservoir of infection for humans and animals. Studies in different geographical areas have reported a prevalence ranging from 5.0% to 39.2% in wild rodents, but most have relied on morphology for identification (Chalmers et al., 1997; Sturdee et al., 1999, 2003; Torres et al., 2000; Bednarska et al., 2003). The limited molecular characterization studies that have been conducted on rodents have identified eight species/genotypes to date: 1. Mouse genotype I, which has been identified in mice and rats in Spain, the UK, Australia and the USA (Morgan et al., 1999b, L. Xiao et al., unpublished) and to date has not been identified in humans. 2. The zoonotic C. parvum, which was detected in mice trapped near sheepgrazing pastures in Victoria, Australia (Morgan et al., 1999b). 3. Cryptosporidium muris, which infects a range of rodents and is generally not zoonotic but which can infect other hosts including dogs, guinea pigs, chipmunks, rabbits, lambs, cats and humans (Iseki et al., 1989; Chalmers et al., 1997; Torres et al., 2000; Hurkova et al., 2003; Hikoska and Nakai, 2005). 4. A novel genotype detected in one wood-mouse (Apodemus sylvaticus) sample from the Czech Republic (Hajdušek et al., 2004). 5. Cryptosporidium meleagridis from a brown rat (Rattus norvegicus) in Japan (Kimura et al., 2007). 6. Eight isolates from brown rats in Japan clustered with sequences denoting W19 and W19 variants found in New York storm water (Jiang et al., 2005; Kimura et al., 2007). 7. Three isolates from brown rats in Japan were identical to a recently described snake isolate 2162 (AY268584) (Xiao et al., 2004b; Kimura et al., 2007). 8. Another isolate from brown rats in Japan showed 100% identity with isolates from non-human primates at Polonnaruwa, Sri Lanka (EF446679) (Ekanayake et al., 2007; Kimura et al., 2007). A recent study in Australia identified mouse genotype I and a novel genotype (mouse genotype II), which is genetically distinct and appears to be hostspecific, as it has not been identified in any other hosts to date (Foo et al., 2007). Unfortunately, it is not possible to determine whether mouse genotype II was the same as the novel mouse genotype described from the Czech Republic, as only the COWP and not the 18S or actin loci were sequenced as part of the Czech study. A previous study, which examined mice from different geographical loci including Australia, the UK and Spain, revealed that the majority of Cryptosporidium-positive mice were infected with mouse genotype I (Morgan et al., 1999b). In that study, five of the Australian mouse samples, which were infected with the zoonotic C. parvum, came from mice which had been trapped on farms in Victoria where large numbers of sheep were grazing. This indicates that sheep may transmit C. parvum to mice, which may in turn transmit Cryptosporidium to other domestic animals. A more recent study by Foo et al. (2007) did not detect the zoonotic C. parvum. This may be because all the positive samples were from arable farms where the mice were unlikely to encounter sheep and cattle. It is possible that mice are only occasionally infected with the C. parvum during periods of heavy environmental contamination.

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Cryptosporidium in Birds Cryptosporidiosis is one of the most prevalent parasitic infections in domesticated, caged and wild birds (Lindsay and Blagburn, 1990; O’Donoghue, 1995), and the parasite has been reported in more than 30 avian species worldwide belonging to the orders Anseriformes, Charadriiformes, Columbiformes, Galliformes, Passeriformes, Psittaciformes and Struthiniformes (Lindsay and Blagburn, 1990; O’Donoghue, 1995; Ng et al., 2006). However, few studies have examined the genetic diversity of Cryptosporidium spp. amongst avian hosts. Currently only three avian Cryptosporidium spp. are recognized: C. meleagridis, C. baileyi and C. galli. These three Cryptosporidium species can each infect a broad range of birds, but they differ in preferred 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, sinuses and trachea. In contrast, C. galli infects only the proventriculus (see Xiao et al., 2004a). 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 that are likely to be re-classified as species in the future, most reports of natural infections have not provided enough information to conclusively determine the species/genotype of Cryptosporidium involved. In addition to the three currently recognized avian species of Cryptosporidium, ten genetically distinct avian genotypes have been described: avian genotypes I–IV from various avian hosts (Meireles et al., 2006; Ng et al., 2006); a novel genotype from a Eurasian woodcock (Scolopax rusticola) (isolate B2-1) from the Czech Republic (Ryan et al., 2003b); a duck genotype from a black duck (Anas rubripes) (Morgan et al., 2001); and goose genotypes I and II, an additional two novel genotypes (geese 3b and 7) from Canada geese (Jellison et al., 2004; Zhou et al., 2004b). Morphometric analysis has not been conducted on most avian genotypes; however, limited morphometric analysis (<10 oocysts per genotype) on avian genotypes II and III suggests that they are morphologically distinct. However, larger numbers of oocysts from each genotype (~50) would need to be analysed in order to produce statistically meaningful results. The results of these recent studies indicate that Cryptosporidium is able to infect a large range of avian species and that, potentially, there are more avian Cryptosporidium species yet to be identified. The potential of these newly identified genotypes to cause disease in birds or humans is unknown, and further studies are required to understand the extent of host adaptation for avian-derived Cryptosporidium genotypes, their species status, transmission dynamics and host range, and the public health implications, if any, of these novel avian genotypes. Four additional species have also been identified in birds: C. hominis, C. parvum, C. muris and C. andersoni (Zhou et al., 2004b; Ng et al., 2006). However, the presence of these oocysts in avian faeces is more likely to be due to mechanical transport rather than an actual infection, as previous studies have shown that, for C. parvum at least, the oocysts were merely passing through the

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digestive tracts of foraging Canada geese without establishing infection (Graczyk et al., 1997).

Cryptosporidium in Fish Little is known about the prevalence or geographical distribution of isolates of Cryptosporidium infecting fish. The first report of Cryptosporidium spp. in fish was in a tropical marine fish (Naso lituratus) in 1981 (Hoover et al., 1981). Cryptosporidium nasorum was subsequently proposed as a species by Levine in 1984 (Levine, 1984). However, only developmental stages of the parasites on the microvillous surface of intestinal epithelial cells were described by light and electron microscopy. No measurements of viable oocysts were provided, and no taxonomically useful diagnostic features were presented. The species was named solely on the basis of the presumed host specificity of Cryptosporidium spp. and is now considered a nomen nudem (Xiao et al., 2004a). Recently, two new piscine Cryptosporidium species have been described: Cryptosporidium molnari from two teleost fish, the gilthead sea bream (Sparus aurata L.) and the European sea bass (Dicentrarchus labrax L.), and Cryptosporidium scophthalmi from cultured turbot (Scophthalmus maximus) (Alvarez-Pellitero and Sitjà-Bobadilla, 2002; Alvarez-Pellitero et al., 2004). Cryptosporidium molnari was found mainly in the stomach epithelium, whereas C. scophthalmi was found mainly in the intestinal epithelium. Oocysts for both species were similar in size to C. parvum and averaged 4.72 × 4.47 mm and 4.44 × 3.91 mm for C. molnari and C. scophthalmi, respectively. In both species, parasite stages were observed deep within the epithelium (Alvarez-Pellitero and Sitjà-Bobadilla, 2002; Alvarez-Pellitero et al., 2004). Unfortunately no molecular characterization of C. molnari or C. scophthalmi has been conducted thus far, and its relationship to other species of Cryptosporidium remains unknown. A recent study conducted histological, genetic and phylogenetic analysis of a C. molnari-like isolate of Cryptosporidium from a guppy (Poecilia reticulata) (Ryan et al., 2004b). The size of the oocysts, the location of the oocysts in the stomach, and the presence of oogonial and sporogonial stages deep within in epithelium, was consistent with C. molnari (Ryan et al., 2004b). Phylogenetic analysis of the 18S rRNA and actin loci revealed that the C. molnari-like isolate formed a distinct group that was basal to all other Cryptosporidium species, suggesting that C. molnari may be the most primitive of the Cryptosporidium spp. However, until genetic sequences are made available for C. molnari and C. scophthalmi, it is impossible to determine the identity of the guppy isolate. Indeed it could be argued that the naming of C. molnari and C. scophthalmi did not follow the criteria for naming species in the genus Cryptosporidium (Xiao et al., 2004a) and therefore should be considered invalid species until genetic and phylogenetic analysis is provided.

Cryptosporidium in Reptiles Cryptosporidium infections are common in reptiles and have been reported in at least 57 reptilian species (O’Donoghue, 1995). Unlike in other animals in which

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Cryptosporidium infection is usually self-limiting in immunocompetent individuals, cryptosporidiosis in reptiles is frequently chronic and sometimes lethal in snakes. Two Cryptosporidium spp. are recognized in reptiles: Cryptosporidium serpentis and C. saurophilum in snakes and lizards, which differ from each other in morphology (oocysts of C. serpentis are bigger than those of C. saurophilum) and predilection sites (C. serpentis is a gastric parasite, whereas C. saurophilum is an intestinal parasite) (Morgan et al., 1999c; Fayer et al., 2000; Xiao et al., 2000, 2004b; Hajdušek et al., 2004). In addition to the two recognized species, a total of seven genotypes have been identified in reptiles including a tortoise genotype, two new snake genotypes, and a lizard genotype, which was genetically distinct but was related to C. serpentis (Xiao et al., 2004b; Alves et al., 2005). Cryptosporidium muris, C. parvum and mouse genotype I have also been identified in reptiles (Morgan et al., 1999c; Xiao et al., 2004b). However, it is likely that that latter three probably do not represent true parasites of these animals but are instead oocysts from rodents ingested by these carnivorous reptiles. This possibility is supported by the fact that the snakes are fed mice and that none of the animals with these oocysts had clinical symptoms. It is also likely that one of the snake genotypes identified by Xiao et al. (2004b) may also be derived from mice, as snake isolate 2162 (AY268584) was identical to Cryptosporidium sequences obtained from brown rats in Japan (Kimura et al., 2007). Currently, there are no effective control strategies against cryptosporidiosis in reptiles. In a small-scale study, it was demonstrated that snakes with clinical and subclinical cryptosporidiosis could be effectively treated with hyperimmune bovine colostrum raised against C. parvum (Graczyk et al., 1998). A common control practice is to euthanize Cryptosporidium-infected snakes, which would prevent the spread of infection to other animals. However one problem with this control measure is the frequent presence of oocysts of C. muris and mouse genotype I in snakes because of the use of feeder mice as part of the diet, which would lead to the killing of uninfected animals.

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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. Veterinary Parasitology 74, 123–132. Guselle, N.J., Appelbee, A.J. and Olson, M.E. (2003) Biology of Cryptosporidium parvum in pigs: from weaning to market. Veterinary Parasitology 113, 7–18. Hajdušek, O., Ditrich, O. and Šlapeta, J. (2004) Molecular identification of Cryptosporidium spp. in animal and human hosts from the Czech Republic. Veterinary Parasitology 122, 183–192. Hamnes, I.S., Gjerde, B.K., Forberg, T. and Robertson, L.J. (2007) Occurrence of Cryptosporidium and Giardia in suckling piglets in Norway. Veterinary Parasitology 144, 222–233. Hikoska, K. and Nakai, Y. (2005) A novel genotype of Cryptosporidium muris from large Japanese field mice, Apodemus speciosus. Parasitology Research 97, 373–379. 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. Journal of Fish Diseases 4, 425–428. Hurkova, L., Hajdušek, O. and Modry, D. (2003) Natural infection of Cryptosporidium muris (Apicomplexa: Cryptosporiidae) in Siberian chipmunks. Journal of Wildlife Diseases 39, 441–444. Iseki, M., Maekawa, T., Moriya, K., Uni, S. and Takada, S. (1989) Infectivity of Cryptosporidium muris (strain RN 66) in various laboratory animals. Parasitology Research 75, 218–222. Izumiyama, S., Furukawa, I., Kuroki, T., Yamai, S., Sugiyama, H., Yagita, K. and Endo, T. (2001) Prevalence of Cryptosporidium parvum infections in weaned piglets and fattening porkers in Kanagawa Prefecture, Japan. Japanese Journal of Infectious Diseases 54, 23–26. 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. Applied and Environmental Microbiology 70, 452–458. Jiang J., Alderisio, K.A. and Xiao, L. (2005) Distribution of Cryptosporidium genotypes in storm event water samples from three watersheds in New York. Applied and Environmental Microbiology 71, 4446–4454. Johnson, J., Buddle, R., Reid, S., Armson, A. and Ryan, U. (2008) Prevalence of Cryptosporidium genotypes in pre- and post-weaned pigs in Australia. Experimental Parasitology 119, 418–421. Kennedy, G.A., Kreitner, G.L. and Strafuss, A.C. (1977) Cryptosporidiosis in three pigs. Journal of the American Veterinary Medical Association 170, 348–350. Kimura, A., Edagawa, A., Okada, K., Takimoto, A., Yonesho, S. and Karanis, P. (2007) Detection and genotyping of Cryptosporidium from brown rats (Rattus norvegicus) captured in an urban area of Japan. Parasitology Research 100, 1417–1420. Langkjær, 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 134, 339–350. Levine, N.D. (1984) Taxonomy and review of the coccidian genus Cryptosporidium (Protozoa, Apicomplexa). Journal of Protozoology 31, 94–98. Lindsay, D.S. and Blagburn, B.L. (1990) Cryptosporidiosis in birds. In: Dubey, J.P., Speer C.A. and Fayer, R. (eds) Cryptosporidiosis in Man and Animals. CRC Press, Boca Raton, FL, pp. 133–148. Maddox-Hyttel, C., Langkjær, R.B., Enemark, H.L. and Vigre, H. (2006) Cryptosporidium and Giardia in different age groups of Danish cattle and pigs: occurrence and management associated risk factors. Veterinary Parasitology 141, 48–55.

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U.M. Ryan and L. Xiao Meireles, M.V., Soares, R.M., dos Santos, M.M. and Gennari, S.M. (2006) Biological studies and molecular characterization of a Cryptosporidium isolate from ostriches (Struthio camelus). Journal of Parasitology 92, 623–626. Mendonça, C., Almeida, A., Castro, A., Delgado, M. de L., Soares, S., da Costa, J.M.C. and Canada, N. (2007) Molecular characterization of Cryptosporidium and Giardia isolates from cattle from Portugal. Veterinary Parasitology 147, 47–50. Morgan, U.M., Constantine, C.C., Forbes, D.A. and Thompson, R.C. (1997) Differentiation between human and animal isolates of Cryptosporidium parvum using rDNA sequencing and direct PCR analysis. Journal of Parasitology 83, 825–830. Morgan, U.M., Buddle, R., Armson, A. Elliot, A. and Thompson, R.C.A. (1999a) Molecular and biological characterisation of Cryptosporidium in pigs. Australian Veterinary Journal 77, 44–47. 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. (1999b) The Cryptosporidium “mouse” genotype is conserved across geographic areas. Journal of Clinical Microbiology 37, 1302–1305. Morgan, U.M., Xiao, L., Fayer, R., Graczyk, T.K., Lal, A.A., Deplazes, P. and Thompson, R.C. (1999c) Phylogenetic analysis of Cryptosporidium isolates from captive reptiles using 18S rDNA sequence data and random amplified polymorphic DNA analysis. Journal of Parasitology 85, 525–530. Morgan, U.M., Monis, P.T., Xiao, L., 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. (2001) Molecular and phylogenetic characterisation of Cryptosporidium from birds. International Journal for Parasitology 31, 289–296. Morgan-Ryan, U.M., Fall, A., Ward, L.A., Hijjawi, N., Sulaiman, I., Fayer, R., Thompson, R.C.A., Olson, M., Lal, A. and Xiao, L.H. (2002) Cryptosporidium hominis n. sp (Apicomplexa: Cryptosporidiidae) from Homo sapiens. Journal of Eukaryotic Microbiology 49, 433–440. Ng, J., Pavlasek, I. and Ryan, U. (2006) Identification of novel Cryptosporidium genotypes from avian hosts. Applied and Environmental Microbiology 72, 7548–7553. O’Donoghue, P.J. (1995) Cryptosporidium and cryptosporidiosis in man and animals. International Journal for Parasitology 25, 139–195. Plutzer, J. and Karanis, P. (2007) Genotype and subtype analyses of Cryptosporidium isolates from cattle in Hungary. Veterinary Parasitology 146, 357–362. 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. Infection Genetics and Evolution 4, 59–66. Power, M.L., Sangster, N.C., Slade, M.B. and Veal, D.A. (2005) Patterns of Cryptosporidium oocyst shedding by eastern grey kangaroos inhabiting an Australian watershed. Applied and Environmental Microbiology 71, 6159–6164. Quílez, J., Sánchez-Acedo, C., Clavel, A., del Cacho, E. and López-Bernad, F. (1996) Prevalence of Cryptosporidium infections in pigs in Aragón (northeastern Spain). Veterinary Parasitology 56, 345–348. Ryan, U.M., Samarasinghe, B., Read, C., Buddle, J.R., Robertson, I.D. and Thompson, R.C.A. (2003a) Identification of a novel Cryptosporidium genotype in pigs. Applied and Environmental Microbiology 69, 3970–3974. Ryan, U.M., Xiao, L., Read, C., Zhou, L., Lal, A.A. and Pavlasek, I. (2003b) Identification of novel Cryptosporidium genotypes from the Czech Republic. Applied and Environmental Microbiology 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.A. and Xiao, L. (2004a) Cryp-

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tosporidium suis n. sp. (Apicomplexa: Cryptosporidiidae) in pigs (Sus scrofa). Journal of Parasitology 90, 769–773. Ryan, U.M., O’Hara, A. and Xiao, L. (2004b) Molecular and biological characterization of a Cryptosporidium molnari-like isolate from a guppy (Poecilia reticulata). Applied and Environmental Microbiology 70, 3761–3765. 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. Applied and Environmental Microbiology 71, 4992–4997. 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. Veterinary Parasitology 122, 103–117. Santín, M., Trout, J.M. and Fayer, R. (2007) Prevalence and molecular characterization of Cryptosporidium and Giardia species and genotypes in sheep in Maryland. Veterinary Parasitology 146, 17–24. Starkey, S.R., Zeigler, P.E., Wade, S.E., Schaaf, S.L. and Mohammed, H.O. (2006) Factors associated with shedding of Cryptosporidium parvum versus Cryptosporidium bovis among dairy cattle in New York State. Journal of the American Veterinary Medical Association 229, 1623–1626. Sturdee, A.P., Chalmers, R.M. and Bull, S.A. (1999) Detection of Cryptosporidium oocysts in wild mammals of mainland Britain. Veterinary Parasitology 80, 273–280. 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. Veterinary Parasitology 116, 97–113. 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. Veterinary Parasitology 92, 253–260. Trout, J.M., Santín, M. and Fayer, R. (2006) Giardia and Cryptosporidium species and genotypes in coyotes (Canis latrans). Journal of Zoo and Wildlife Medicine 37, 141–144. Vitovec, J., Hamadejova, K., Landova, L., Kvac, M., Kvetonova, D. and Sak, B. (2006) Prevalence and pathogenicity of Cryptosporidium suis in pre- and post-weaned pigs. Journal of Veterinary Medicine Series B 53, 239–243. Warren, K.S., Swan, R.A., Morgan-Ryan, U.M., Friend, J.A. and Elliot, A. (2003) Cryptosporidium muris infection in bilbies (Macrotis lagotis). Australian Veterinary Journal 81, 739–741. Wieler, L.H., Ilieff, A., Herbst, W., Bauer, C., Vieler, E., Bauerfeind, R., Failing, K., Klos, H., Wengert, D., Baljer, G. and Zahner, H. (2001) Prevalence of enteropathogens in suckling and weaned piglets with diarrhoea in southern Germany. Journal of Veterinary Medicine Series B 48, 151–159. 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. Applied and Environmental Microbiology 65, 1578–1583. 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. Applied and Environmental Microbiology 65, 3386–3391. Xiao, L., Morgan, U.M., Fayer, R., Thompson, R.C. and Lal, A.A. (2000) Cryptosporidium systematics and implications for public health. Parasitology Today 16, 287–292. 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. Journal of Infectious Diseases 185, 1846–1848.

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Insights Into the Molecular Detection of Giardia duodenalis: Implications for Epidemiology S.M. CACCIÒ1, M. LALLE1, R. BECK2 AND E. POZIO1 1Istituto

Superiore di Sanità, Rome, Italy; 2University of Zagreb, Croatia

Abstract Giardia duodenalis is a widespread parasite of mammalian species, including humans. Due to its invariant morphology, investigation of aspects such as host specificity and transmission patterns requires a direct genetic characterization of cysts/trophozoites from host samples. A number of molecular assays have been developed to help in unravelling the complex epidemiology of this infection. Recently, however, molecular approaches have been complicated by the recognition of intra-isolate sequence heterogeneity (i.e. ‘mixed templates’), which affects subtype identification and the assignment of isolates to specific G. duodenalis assemblages. This raises concerns about the previous interpretation of genotyping data, and indicates the need to understand the mechanisms that are responsible for these unexpected findings. In this chapter, we critically review those different mechanisms and discuss some possible experimental strategies that can be used in future studies.

Introduction Giardia is a genus of intestinal flagellates that infect a wide range of vertebrate hosts. The genus currently comprises six species; namely Giardia agilis detected in amphibians, Giardia ardeae and Giardia psittaci detected in birds, Giardia microti and Giardia muris detected in rodents, and Giardia duodenalis detected in a wide spectrum of mammals, which are distinguished on the basis of the morphology and ultrastructure of their trophozoites (Adam, 2000). Giardia duodenalis (syn. G. intestinalis, G. lamblia) is the only species found in humans, although it is also found in other mammals, including domestic pets and livestock (Thompson, 2004). A considerable amount of data have shown that G. duodenalis should be considered as a species complex whose members, while showing little variation in their morphology, can be assigned to at least seven distinct assemblages (A to G) based on genetic analyses (Monis et al., 2003). Only assemblages A and B have been detected in humans and in a wide © CAB International 2009. Giardia and Cryptosporidium: From Molecules to Disease (eds G. Ortega-Pierres et al.)

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Giardia duodenalis assemblages and their distribution in mammalian

Assemblages

Hosts

A

Humans and other primates, livestock, horses, dogs, cats, guinea pigs, fallow deer, white-tailed deer, ferrets Humans and other primates, livestock, horses, dogs, coyotes, muskrats, beavers Dogs, cats, coyotes, wolves Cattle, sheep, goats, water buffaloes, muflons Cats Rats

B C, D E F G

range of other mammalian hosts, whereas the remaining assemblages (C to G) are likely to be host-specific (Table 7.1). Due to the lack of variation in cyst and trophozoite morphology, the reliable discrimination of G. duodenalis isolates is necessarily based on their genetic characterization. Over the last decade or so, specially after the introduction of DNA amplification techniques, this approach has been extensively used, in particular to understand the role of animals in the epidemiology of human infection, i.e. to estimate the extent of zoonotic transmission (Cacciò et al., 2005), as well as to detect and genotype Giardia spp. cysts from water and food samples (Smith et al., 2006). Most of these studies, however, were based on the analysis of single markers, and no systematic evaluation of the genetic variability and usefulness of the different genetic loci has been performed. Indeed, the recognition of subgroups within assemblages A and B, as well as the extent of genetic variability among G. duodenalis isolates, are mainly based on isoenzyme analysis (Monis et al., 2003), an approach which is not routinely applicable to clinical and environmental samples, as it requires the in vitro propagation of Giardia spp. cysts.

Current Genotyping Approach Molecular typing: genetic loci, their variability, and the issue of nomenclature Compared to other protozoan pathogens, the genotyping technique for Giardia spp. is not particularly advanced, and the vast majority of studies rely on the analysis of the small subunit ribosomal RNA (ssu-rRNA), the β-giardin (bg), glutamate dehydrogenase (gdh), elongation factor 1-alpha (ef-1), triose phosphate isomerase (tpi), and the GLORF-C4 (C4) genes (reviewed in Cacciò et al., 2005). Some of these genes have been mapped, using hybridization on chromosomes separated by pulsed field gel electrophoresis (Adam, 2000), on to chromosome 5 (tpi and bg genes), chromosome 4 (gdh gene) and chromosome 1 (the majority of the ssu-rRNA gene copies). A search of the latest (April 2007) version of the G. duodenalis database (see http://gmod.mbl.edu/) showed that the tpi and bg genes are located in supercontig 577 (2 mega-bases (Mb) in size), and are

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about 150 kb apart. Similarly, the ef-1 gene is located on supercontig 608 (64 kb), and the C4 gene is located on supercontig 573 (1.5 Mb). Therefore, the genes can be considered unlinked in the G. duodenalis genome. This is an important and desirable feature for genotyping studies. A large number of sequences have been obtained from each of these genes and from isolates of human and animal origin collected worldwide (120 for ssu-rRNA, 170 for bg, 134 for gdh, 10 for ef-1, 70 for tpi, and 3 for C4, according to a search of the April 2007 GenBank release). Unfortunately, most sequences are partial, particularly in the case of the ssu-rRNA gene and, in general there is a tendency to reduce the amplicon size, either to increase PCR sensitivity or as a result of the design of new primers when more sequences become available. In terms of their polymorphism, these genes are quite different, with tpi and gdh genes being the more variable, followed by bg and C4 genes, then by the more conserved ef-1 and ssu-rRNA genes (Monis et al., 1999). This is also reflected by differences in substitution patterns, with the bg and ef-1 genes showing few, if any, non-synonymous changes, whereas the tpi and gdh genes appear to also tolerate amino acid replacements. Previous analyses, mainly using G. duodenalis isolates adapted to in vitro growth, and based on isoenzymes (Meloni et al., 1995) and DNA sequence analyses (Monis et al., 1999), have shown the existence of two subgroups in assemblage A (AI and AII) and in assemblage B (BIII and BIV). It is clear, however, that more variability is present in these as well as in isolates from the other G. duodenalis assemblages (Monis et al., 2003). Substructuring was evident within this assemblage, with isolates from the same host species forming distinct subclusters, which may correspond to yet undefined subgroups (Monis et al., 2003). More recent DNA-based genotyping studies have indeed identified a number of subtypes from both human and animal isolates (e.g. Sulaiman et al., 2003; Lalle et al., 2005a) and have confirmed substructuring within major G. duodenalis assemblages (Fig. 7.1). In these studies, a different terminology was used to describe subtypes, such as A3, A4, A5, or B1, B2, B3 at the bg locus (Lalle et al., 2005b), BS1, BS2, BS3, or CS1, CS2, CS3 at the tpi locus, or even B0, B1, B2 at the gdh locus. This has generated a certain degree of confusion, particularly when different subtypes were found by sequencing different genetic markers, i.e. when there was a lack of concurrence between the results for these genes (Robertson et al., 2006).

Epidemiology of Giardiasis Several characteristics of G. duodenalis influence the epidemiology of infection: ● ●

●

●

The infective dose is low (about 10–100 cysts; Rendtorff, 1954). Cysts are immediately infectious when excreted in faeces, and can be transmitted by person-to person or animal-to-animal contact (Thompson, 2004). Cysts are remarkably stable and can survive for weeks to months in the environment (Smith et al., 2006). Environmental contamination can lead to the contamination of drinking water and food (Karanis et al., 2007).

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98 99

AI A II A

100

F (cat) 100

E (hoofed animals)

91 100

B III

G. duodenalis

100 B IV

84

100 100

D (dog)

C (dog)

G (rat) 100 G. microti

G. ardeae G. muris

0.05

Fig. 7.1. Phylogenetic relationships of Giardia spp. inferred by the neighbour-joining analysis of the triose phosphate isomerase (tpi) nucleotide sequences. Only bootstrap values >70 are indicated. Modified from Lalle et al. (2007).

Direct and indirect transmission between infected hosts and susceptible individuals is favoured by high population densities (e.g. during lambing or calving), and by close contact (e.g. during recreational bathing or consumption of contaminated water) (Cacciò et al., 2005). Human isolates from different geographical locations, examined by PCR amplification of DNA extracted directly from faeces, demonstrate that only G. duodenalis assemblages A and B are associated with human infections. The prevalence of each assemblage varies considerably from country to country; assemblage B seems more common overall, but no strong conclusions can be drawn from current data (Cacciò et al., 2005). One important and still debated aspect in the epidemiology of human giardiasis is the extent of zoonotic transmission (Thompson, 2004). A number of animals have been considered to act as reservoirs, including livestock, pets and aquatic animals, but although the World Health Organization has considered G. duodenalis to have zoonotic potential for over 20 years (WHO, 1979), direct evidence is still lacking. As an example, beavers are often indicated as a source

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of environmental contamination and potential waterborne infection in humans (giardiasis is known as ‘beaver fever’ in Canada), but application of molecular techniques to samples collected in British Columbia (from humans, beavers and drinking water) could not support a causative link (McIntyre et al., 2000). Furthermore, the isoenzyme data of Monis et al. (2003) have shown that human and animal-derived isolates of G. duodenalis from assemblages A and B are rarely identical, and tend to form separate clusters in phylogenetic analyses. Finally, typing of wildlife isolates worldwide increasingly shows a predominance of hostadapted genotypes. Isolates from wild hoofed animals are often typed as assemblage A, but they are so genetically distinct from human isolates that zoonotic transmission seems unlikely (Lalle et al., 2007). Therefore, the available data, albeit still fragmentary, indicate a minor role of animals in the epidemiology of human giardiasis. However, zoonotic transmission may occur in endemic foci, where humans and animals live in close association with each other. In such cases, molecular characterization of cysts and detailed epidemiological investigations can provide insights into transmission patterns, as recently shown by a study of giardiasis in humans and dogs from a remote tea-growing community of north-east India (Traub et al., 2004). Certainly, we need more sensitive molecular methods to assess the zoonotic potential of Giardia spp. cysts shed by animals. In this respect, an important lesson can be learned from studies on cryptosporidiosis. Here, human infection with Cryptosporidium parvum has been traditionally considered zoonotic, with cattle being the main reservoir (Cacciò, 2005). However, the application of fingerprinting techniques, based on a combination of highly polymorphic micro- and minisatellite markers, has shown that a number of C. parvum types appear restricted to humans and may be maintained largely, if not exclusively, by an anthroponotic cycle (Mallon et al., 2003). Whether a similar situation exists for giardiasis is not known, but deserves to be considered. Transmission of G. duodenalis through drinking and recreational water is well documented, as are outbreaks of giardiasis following the consumption of contaminated water. Indeed, of the 325 water-associated outbreaks of parasitic protozoan disease reported so far (Karanis et al., 2007), G. duodenalis was the causative agent in about 40%. Despite this fact, little is known about the Giardia spp. or G. duodenalis assemblages present in the aquatic environment (Smith et al., 2006). In all wastewater samples examined, only G. duodenalis cysts were identified, and, more importantly, only human pathogens (assemblages A and B) were found (Cacciò et al., 2003; Sulaiman et al., 2004). This is in sharp contrast with results obtained for Cryptosporidium spp., but probably reflects the nature of the samples tested (raw urban wastewater) and the very limited number of surveys conducted. Most foodborne outbreaks of giardiasis were ascribed to direct contamination by a food handler, but in two instances a role for zoonotic transmission was suggested, namely the consumption of a Christmas pudding contaminated with rodent faeces and tripe soup made from the offal of an infected sheep (Smith et al., 2006). Notably, no information is available on the G. duodenalis assemblage(s) associated with these outbreaks.

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Current Problems in Molecular Genotyping Mixed templates: impact and possible origin The presence of mixed templates, characterized by two overlapping nucleotide peaks at specific positions in the electropherograms, was already reported 10 years ago (Hopkins et al., 1997). As a matter of fact, a number of sequences submitted to GenBank (for example, U47632) over the last few years contained undetermined positions (R, Y, etc.) and are likely to represent cases of mixed templates. The impact of this phenomenon is quite evident, as it hampers the precise assignment of isolates at the assemblage or subtype level. This is particularly relevant for assemblage B isolates, where a considerable fraction of isolates are characterized by mixed templates at different genetic loci (unpublished data from our laboratory). Two principal mechanisms can explain the occurrence of mixed templates: (i) ‘true’ mixed infections; and (ii) allelic sequence heterozygosity (ASH).

Mixed infections The occurrence of mixed infections has been reported in molecular-based surveys performed in Australia, the UK, India, Italy and Ethiopia (Hopkins et al., 1997; Amar et al., 2002; Traub et al., 2004; Lalle et al., 2005a, 2005b; Gelanew et al., 2007). The percentage of mixed infections ranged from 2.0% to 21.0% and was higher in less economically developed countries. Mixed infections are known to occur both at the inter- and intra-assemblage levels (for example, A plus B, or AI plus AII). In such cases, co-amplification of genetic material from genetically different cysts, followed by direct sequencing of PCR products, may result in heterogeneous sequencing profiles.

Allelic sequence heterozygosity Allelic sequence heterozygosity (ASH) has long been considered to be an explanation for mixed templates. Baruch et al. (1996) first reported ASH by sequence analysis of the tpi gene. They sequenced multiple cloned PCR products to demonstrate that the ambiguity in sequencing gels was due to ASH and not to technical artefacts. Interestingly, ASH was found between two human isolates from the AII subgroup, as well as between assemblage B isolates of human and animal origin. The level of ASH, however, was extremely low, less than 0.1%, a finding that seems to be confirmed by the genome sequencing project. However, the assembled portion of the genome does not include sub-telomeric regions, where chromosome size heterogeneity (heteromorphy) is known to occur (Adam, 2000). These regions carry copies of the rDNA of variable surface protein genes, and also of retroposons (Prabhu et al., 2007). Therefore, G. duodenalis

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chromosomes appear to be organized into conserved, central domains and polymorphic chromosome ends, resembling other parasitic genomes (Lanzer et al., 1995). A low ASH level is considered surprising for a polyploid, asexually replicating organism such as G. duodenalis (Adam, 2000). Indeed, substantial differences are expected to accumulate between the chromosome homologues in asexual organisms with a ploidy of two or higher, as has been shown for bdelloid rotifers (Welch and Meselson, 2000). In this context, it is important to note that in other asexual organisms, such as Daphnia spp., the rate of loss of nucleotide heterozygosity by ameiotic recombination is substantially greater than the rate of introduction of new variation by mutation. This suggests that the evolutionary potential of asexual diploid species is not only a matter of mutation accumulation and reduced efficiency of selection, but it underscores the limited utility of using neutral allelic divergence as an indicator of ancient asexuality (Omilian et al., 2006). Recent studies have shown that each nucleus has at least one copy of the genome, and that the two nuclei are partitioned equationally at cytokinesis (Yu et al., 2002), therefore ruling out two possible explanations for low ASH, namely that each nucleus contains a different genome, or that nuclear segregation is reductional (Yu et al., 2002). This result, however, was not confirmed by another study (Tumova et al., 2006), which reported differences in the number and size of chromosomes between the nuclei of single cells and, notably, suggested that each nucleus behaves as a clonal lineage, and should therefore accumulate mutations independently. Clearly, this complex issue deserves future investigation.

Assemblage swapping: impact and possible origin Recent publications (Read et al., 2004; Traub et al., 2004; Gelanew et al., 2007) have reported that the assignment of isolates to specific G. duodenalis assemblages is not always reliable, as different markers can give different results. This has been found both in human and animal isolates, appears to be rather frequent (up to 25% of dog isolates; Read et al., 2004), and has been found using different combinations of gene markers. It is of particular relevance that animal isolates can be typed as ‘potentially zoonotic’ with one marker, but as ‘host-specific’ with another. For molecular epidemiological studies, therefore, this has very important implications, as totally different conclusions may be drawn, depending on the way that genotyping data are obtained and interpreted. A number of mechanisms have been proposed to account for assemblage swapping, including preferential amplification of one assemblage over another (in cases of mixed infection), introgression, retention of ancestral polymorphisms and, more generally, recombinational (sexual) events. The occurrence of mixed infections has been summarized above. The possibility that a high percentage of infections are sustained by two genetically different parasites cannot be disregarded, particularly in regions of high endemicity.

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Introgression Introgression requires hybridization between closely related species and backcrossing between hybrids and one of the parental populations to produce gene flow (Anderson, 2001). This mechanism can result in segregation of divergent alleles in the same population. Conversely, identical alleles can occur in genetically distinct populations due to the retention of ancestral polymorphism, if the time of divergence of sister taxa from a common ancestor has been insufficient to allow fixation of the alleles produced by mutation and genetic drift (Anderson, 2001). If the organisms under study are morphologically identical, as is the case for G. duodenalis, the use of single-locus markers in molecular epidemiological studies may lead to ambiguous or even wrong conclusions, as demonstrated by Anderson (2001) using Ascaris spp. as a case study.

Meiosis and sex The evidence for the presence of sexuality in flagellates of the genus Giardia is, at present, only indirect. Using a phylogenomic approach, Ramesh et al. (2005) identified a core group of meiotic genes in the G. duodenalis genome (that are true orthologues of their counterparts in sexual organisms) and concluded that the parasite has the potential to perform meiosis, and thus, sexual reproduction. In a study of genome ploidy in different life cycle stages (Bernander et al., 2001), it has been shown that, during excystation, the recently excysted cell divides twice without DNA replication. The authors commented that: differentiation in G. lamblia is therefore reminiscent of meiosis, in which the genome is first replicated and then divided twice without DNA replication. It is possible that differentiation of primitive eukaryotes into cystic forms is an ancestral form of sexual processes.

If meiosis is present in G. duodenalis, what we need to know is how the diploidy is restored (Birky, 2005), and if this happens between nuclei from two different individuals (outcrossing) or from the same individual (selfing), as the net result of these two alternatives is crucially different.

Clonality As mentioned above, other studies have challenged this view, and suggested a ‘clonal’ mode of reproduction (Tumova et al., 2006). The (apparent) stable transmission of an aneuploid pattern is possible only if meiosis is absent, as the presence of homologous copies of each chromosome is a prerequisite for the first meiotic division (Tumova et al., 2006). In the early 1990s, Tibayrenc et al. (1990) proposed a clonal theory of population structure where sexual recombination is not frequent enough to break the prevalent pattern of clonal population structure. In this study, allozyme data were used to propose that G. duodenalis is a clonally propagating organism.

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The concept of clonality was then challenged by Maynard-Smith et al. (1993), who proposed an epidemic population structure where there is a background level of frequent sexual recombination with occasional clonal expansion of a few particular genotypes. Whether the population structure of G. duodenalis is clonal, panmictic or epidemic remains an open question, mainly because an insufficient number of isolates have been tested, and statistical tests could not be applied (Monis et al., 2003). Considering the conceptual and practical implications of clonality, this is clearly an important area for future investigation (Tibayrenc, 2005).

Mobile Elements and the Giardia-specific Virus Other mechanisms that could enhance genetic exchanges within and between trophozoites should also be considered. In particular, genetic mobile elements have been identified in the G. duodenalis genome, and a Giardia-specific virus has been widely studied. Giardia duodenalis harbours non-LTR (non-long terminal repeats) retroelements of the LINEs family (long interspersed nuclear elements), that are transposed by reverse transcription of mRNA directly into the site of integration. Most G. duodenalis sub-telomeres consist of tandem copies of active LINE retroposons (either GilM or GilT elements), which directly abut the telomeric repeats and are oriented such that reverse transcription would have run towards the chromosome end (Wickstead et al., 2003). Repeats play an integral part in ongoing genomic evolution and can perform diverse roles at different times, imparting a greater changeability to genomes. When an organism faces a changeable environment, the advantages of genomic flexibility (particularly if it can be contained at specific loci) may outweigh the extra cost of replication and of mutagenic effect exerted on other genomic regions. It is currently unknown what role the retroposons are playing (or have played) in the evolution of the G. duodenalis genome. Giardia-specific virus (GLV) is a double-stranded (ds) RNA virus of the Totiviridae family, constituted by a 36 nm non-enveloped icosahedron comprising one dsRNA of about 7 kb (Wang and Wang, 1991). GLV infects many G. duodenalis from assemblages A, B, C/D and E, although no correlation between the presence or absence of the virus and the specific assemblage has been found (Sedinova et al., 2003; Chen et al., 2007), and cohabitation of multiple GLV species in the same parasite has also been demonstrated (Tai et al., 1996). In G. duodenalis Portland I strain, which is chronically infected by this virus, viral RNA was detected in the cytoplasm as well as in the twin nuclei (Tai et al., 1991). Recombinant GLV cDNA has been successfully introduced into GLV-infected trophozoites to express a heterologous gene in G. duodenalis. Moreover, the chimeric RNA could be replicated as double-stranded RNA and packaged into virus-like particles, and the recombinant virions, by themselves, can superinfect G. duodenalis trophozoites and start new rounds of expression (Yu et al., 1996). All these observations are compatible with fragment(s) of the G. duodenalis genome being inserted in the GLV and then shuttled between trophozoites of different G. duodenalis assemblages.

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Intriguingly, plasmid DNA transfected into trophozoites of assemblage A is maintained as a multimeric episome, whereas in assemblage B it is always integrated into the genome by homologous recombination, with insertion of multiple copies (Singer et al., 1998). This contrasting behaviour could reflect isolate-specific differences in the factors determining whether exogenous DNA is integrated or maintained episomally. If plasmid- or virus-mediated lateral exchange of DNA between two trophozoites can occur, homologous recombination could, at least partially, explain the higher rate of ASH observed in assemblage B compared with assemblage A.

Future Experimental Strategies To understand the origin of mixed templates, different approaches can be envisaged, including the use of assemblage-specific primers (to estimate more precisely the occurrence of mixed infections and its contribution to both phenomena), the cloning of PCR products from single isolates followed by sequencing of multiple clones (to evaluate ASH), and the direct typing of single nuclei isolated from cultured trophozoites (to prove genetic heterogeneity between the two nuclei). For a more comprehensive analysis, however, it is still invaluable to obtain and maintain cloned isolates that are representative of each assemblage. This research will also benefit from international collaboration and from the use of validated sets of markers and methods that allow direct comparisons between studies.

References Adam, R.D. (2000) The Giardia lamblia genome. International Journal for Parasitology 30, 475–484. Amar, C.F.L., Dear, P.H., Pedraza-Díaz, S., Looker, N., Linnane, E. and McLauchlin, J. (2002) Sensitive PCR-restriction fragment length polymorphism assay for detection and genotyping of Giardia duodenalis in human feces. Journal of Clinical Microbiology 40, 446–452. Anderson, T.J.C. (2001) The dangers of using single locus markers in parasite epidemiology: Ascaris as a case study. Trends in Parasitology 17, 183–188. Baruch, A.C., Isaac-Renton, J. and Adam, R.D. (1996) The molecular epidemiology of Giardia lamblia: a sequence-based approach. Journal of Infectious Diseases 174, 233–236. Bernander, R., Palm, J.E. and Svard, S.G. (2001) Genome ploidy in different stages of the Giardia lamblia life cycle. Cellular Microbiology 3, 55–62. Birky, C.W., Jr (2005) Sex: is Giardia doing it in the dark? Current Biology 15, 56–58. Cacciò, S.M. (2005) Molecular epidemiology of human cryptosporidiosis. Parassitologia 47, 185–192. Cacciò, S.M., De Giacomo, M., Aulicino, F. and Pozio, E. (2003) Giardia cysts in wastewater treatment plants in Italy. Applied and Environmental Microbiology 69, 3393–3398. Cacciò, S.M., Thompson, R.C.A., McLauchlin, J. and Smith, H.V. (2005) Unravelling Cryptosporidium and Giardia epidemiology. Trends in Parasitology 21, 430–437.

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Chen, L., Li, J., Zhang, X., Liu, Q., Yin, J., Yao, L., Zhao, Y. and Cao, L. (2007) Inhibition of krr1 gene expression in Giardia canis by a virus-mediated hammerhead ribozyme. Veterinary Parasitology 143, 14–20. Gelanew, T., Lalle, M., Hailu, A., Pozio, E. and Cacciò, S.M. (2007) Molecular characterization of human isolates of Giardia duodenalis from Ethiopia. Acta Tropica 102, 92–99. Hopkins, R.M., Meloni, B.P., Groth, D.M., Wetherall, J.D., Reynoldson, J.A. and Thompson, R.C. (1997) Ribosomal RNA sequencing reveals differences between the genotypes of Giardia isolates recovered from humans and dogs living in the same locality. Journal of Parasitology 83, 44–51. Karanis, P., Kourenti, C. and Smith, H. (2007) Waterborne transmission of protozoan parasites: a worldwide review of outbreaks and lessons learnt. Journal of Water Health 5, 1–38. Lalle, M., Jimenez, E., Cacciò, S.M. and Pozio, E. (2005a) Genotyping of Giardia duodenalis from humans and dogs from Mexico using β-giardin nested PCR assay. Journal of Parasitology 91, 203–205. Lalle, M., Pozio, E., Capelli, G., Bruschi, F., Crotti, D. and Cacciò, S.M. (2005b) Genetic heterogeneity at the b-giardin locus among human and animal isolates of Giardia duodenalis and identification of potentially zoonotic subgenotypes. International Journal for Parasitology 35, 207–213. Lalle, M., Frangipane di Regalbono, A., Poppi, L., Nobili, G., Tonanzi, D., Pozio, E. and Cacciò, S.M. (2007) A novel Giardia duodenalis assemblage A subtype in fallow deer. Journal of Parasitology 93, 426–428. Lanzer, M., Fischer, K. and Le Blancq, S.M. (1995) Parasitism and chromosome dynamics in protozoan parasites: is there a connection? Molecular and Biochemical Parasitology 70, 1–8. Mallon, M.E., MacLeod, A., Wastling, J.M., Smith, H. and Tait, A. (2003) Multilocus genotyping of Cryptosporidium parvum Type 2: population genetics and sub-structuring. Infection Genetics and Evolution 3, 207–218. Maynard-Smith, J., Smith, N.H., O’Rourke, M. and Spratt, B.G. (1993) How clonal are bacteria? Proceedings of the National Academy of Sciences of the USA 90, 4384–4388. McIntyre, L., Hoang, L., Ong, C.S., Lee, P. and Isaac-Renton, J.L. (2000) Evaluation of molecular techniques to biotype Giardia duodenalis collected during an outbreak. Journal of Parasitology 86, 172–177. Meloni, B.P., Lymbery, A.J. and Thompson, R.C.A. (1995) Genetic characterization of isolates of Giardia duodenalis by enzyme electrophoresis: implications for reproductive biology, population structure, taxonomy, and epidemiology. Journal of Parasitology 81, 368–383. Monis, P.T., Andrews, R.H., Mayrhofer, G. and Ey, P.L. (1999) Molecular systematics of the parasitic protozoan Giardia intestinalis. Molecular Biology and Evolution 16, 1135–1144. Monis, P.T., Andrews, R.H., Mayrhofer, G. and Ey, P.L. (2003) Genetic diversity within the morphological species Giardia intestinalis and its relationship to host origin. Infection Genetics and Evolution 3, 29–38. Omilian, A.R., Cristescu, M.E., Dudycha, J.L. and Lynch, M. (2006) Ameiotic recombination in asexual lineages of Daphnia. Proceedings of the National Academy of Sciences of the USA 103, 18638–18643. Prabhu, A., Morrison, H.G., Martinez, C.R., III and Adam, R.D. (2007) Characterisation of the subtelomeric regions of Giardia lamblia genome isolate WBC6. International Journal for Parasitology 37, 503–513.

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Wildlife with Giardia: Villain, or Victim and Vector? S.J. KUTZ1, R.C.A. THOMPSON2 AND L. POLLEY3 1Faculty

of Veterinary Medicine, University of Calgary, Alberta, Canada; Collaborating Centre for the Molecular Epidemiology of Parasitic Infections, Murdoch University, WA, Australia; 3Western College of Veterinary Medicine, University of Saskatchewan, Canada 2WHO

Abstract Among mammalian wildlife, Giardia has been reported on every continent and from a wide variety of species, including marsupials, rodents, insectivores, ungulates, marine mammals, felids, canids and ursids. Increasing efforts have been made to determine the strain and zoonotic potential of Giardia and this has resulted in new host and geographical records for Giardia duodenalis. According to many peer-reviewed publications, the mere presence of Giardia in wildlife is sufficient to implicate wildlife as source of infection for humans. However, the bulk of the evidence to support this claim demonstrates correlation, not causation. In fact, in many cases wildlife are more likely to be the victims, with ‘spillover’ of zoonotic strains from people and/or domestic animals. A critical re-evaluation of zoonotic strains of Giardia in wildlife is needed. This should include a return to classical parasitology and epidemiology in order to develop a quantitative understanding of the temporal and spatial patterns and impacts of the various Giardia strains in wildlife and the potential interactions with domestic systems. It has become abundantly clear that the mere detection of Giardia cysts in wildlife is no longer sufficient evidence of zoonotic transmission and that there remains considerable uncertainty about the structure of the parasite web and the direction and intensity of parasite flow for ‘zoonotic’ strains of Giardia among domestic and wild animal, and human hosts.

Introduction Since its discovery in 1681, Giardia has been recovered from a wide range of mammalian, avian, reptilian, and amphibian hosts, many of which have been considered to be important reservoirs of infection for people (Thompson, 2002). At one time over 50 species of Giardia had been described. The recent application of molecular tools has led to a rapid evolution in our understanding of the parasite’s taxonomy and host specificity, confirming some previously named species while refuting others. Importantly, recognition of these host specificities has generated doubt concerning the ubiquitous zoonotic threat once thought 94

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to be associated with Giardia. Today there are seven recognized species assemblages defined on the basis of molecular and phylogenetic analyses. Based on our current understanding, only Giardia duodenalis assemblages A and B, described from a variety of mammalian species, have demonstrated zoonotic potential, although the zoonotic potential of novel subtypes is unclear.

Are Wildlife Species Significant Reservoirs for Zoonotic Transmission of Giardia? Parasites of the genus Giardia are widespread in mammalian, avian, reptilian and amphibian wildlife. Mammalian hosts, in particular, have long been considered important sources of infection for people. In fact, the beaver is considered such a common source of human infection that in North America the disease in people is often referred to as ‘beaver fever’. The link to beavers is so ingrained among the public and healthcare professionals that physicians have been known to advise that there is no risk of Giardia in areas where beavers are absent, and one peer-reviewed paper suggested that the movement of beavers northwards will enhance the emergence of giardiasis in people in northern regions (Parkinson and Butler, 2005). The belief in the integral role of beavers in the propagation of giardiasis is also reflected in the active education campaigns in parks and protected areas across Canada and the USA, in the culture of campers and backcountry hikers, and is a tremendous stimulus for the purchase of water filters. While several wildlife species are, to varying degrees, suitable hosts for the human Giardia duodenalis assemblages A and B, beavers appear to be particularly susceptible to these strains, are able to maintain them for several months and, once infected, may serve as suitable reservoirs, and perhaps amplifiers, for spill-back to people (Davies and Hibler, 1979). Most of the evidence implicating beavers and other wildlife hosts as significant sources of Giardia for people is, however, anecdotal and correlational – direct links have rarely, if ever, been established (Appelbee et al., 2002; Sulaiman et al., 2003; Cacciò et al., 2005). One of the most extensive epidemiological studies examining the links between wildlife, domestic animals and people, is that by Davies and Hibler from 1975 to 1977 (Davies and Hibler, 1979). These authors investigated Giardia in multiple species (26 wildlife, 5 domestic – dogs, cats, cattle, horses, sheep – and people) in multiple watersheds in Colorado, USA. They also completed a series of experimental cross-infections using Giardia from different host species to determine host specificity. This landmark study demonstrated several important aspects of the spatial, temporal, and host distribution of Giardia: ● ●

●

Giardia was already present in people and dogs entering wilderness areas. Of 26 wildlife species sampled, only beavers and coyotes were infected with Giardia in the wild. There was a temporal pattern of Giardia infection for beavers, coyotes and cattle, with none of these species infected in the spring but all infected by autumn, suggesting that the parasite did not overwinter in these species, but that they became infected during the summer.

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●

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Prevalence in beavers was higher at or downstream from outbreak locations or human settlements, campgrounds, or sewage plants, compared with upstream, and prevalence in beavers increased with distance downstream. Giardia from people, beavers or (captive) deer varied in their ability to infect other host species.

Through this survey, Davies and Hibler established host, temporal and spatial distributions of Giardia spp. in areas of Colorado, and through their experimental work they established pre-patent and patent periods and reported any clinical signs associated with the parasite in various host species, including people (Davies and Hibler, 1979). They also observed that ‘beaver may be an excellent sentry animal’ for watershed contamination, and that once infected ‘beaver are an important reservoir … they carry the infection for at least 3 months and defecate into the water where Giardia cysts survive very well’. Domestic livestock were considered as part of the transmission cycle in that they congregate around waterbodies and defecate in and around them and thus may become important reservoirs. Dogs were also considered as important sources of infection and disseminators of Giardia because they accompany people everywhere, defecate at will, are coprophagic, and often roll in faeces. Ultimately Davies and Hibler (1979) stated that ‘Humans are the most important component in the epidemiology of giardiasis’. They commented that: people use watersheds heavily and frequently defecate indiscriminately in or near waterbodies (the ‘wilderness experience’); ‘human feces and toilet paper were found in several locations’ along waterbodies; used diapers have been found in lakes and streams, and; there are several reports of people emptying recreational vehicle toilets into rivers. They concluded that ‘giardiasis is a growing problem and will remain so for many years’ because there will be increased use of wilderness areas by people and their pets (some of which bring Giardia with them), and there is a lack of sanitary facilities and a reluctance of people to use them (Davies and Hibler 1979). Subsequent advances in molecular biology have allowed us to unravel some of the issues associated with host-specificity, and the zoonotic potential of some strains of Giardia has been confirmed. Despite these developments, no studies have demonstrated unequivocally that wildlife serve as the origin or main reservoir for human infections with the parasite. In many waterborne outbreaks affecting people, sewage contamination of the water supply has been implicated as the source of Giardia (Craun et al., 2005). In reports where wildlife has been identified as the source of the parasite, supporting evidence is lacking and wildlife appear to be blamed by default (Charron et al., 2004). Additionally, Giardia cysts are invariably much more abundant in sewage outflow (treated and untreated) than in the raw water intake in most communities, suggesting that contamination of waterbodies is of human (or domestic animal) origin. It seems that beavers, and other wildlife using water downstream from these outflows, are unfortunate victims of this environmental contamination (Roach et al., 1993; Craun et al., 2005). The uncertainty of the relationship between Giardia in wildlife and people was eloquently captured by Cacciò et al. (2005):

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The occurrence of Giardia in wildlife … has been the single most important factor implicating Giardia as a zoonotic agent. It is therefore surprising that there is little evidence to support the role of wildlife as a source of disease in humans. Although wildlife, particularly aquatic mammals, is commonly infected with Giardia, there is little evidence to implicate such infections as the original contaminating source in waterborne outbreaks. It would seem that such animals are more likely to have become infected from water contaminated with faecal material of human or, less likely, domestic animal origin, thus serving to amplify the numbers of the originally contaminating isolate. The few studies that have genotyped Giardia of beaver origin have confirmed previous suggestions that the source of Giardia infection in beavers was likely to be of human origin.

It has become abundantly clear that the mere detection of Giardia cysts in wildlife is no longer sufficient evidence of zoonotic transmission and that there remains considerable uncertainty about the structure of the parasite web and the direction and intensity of parasite flow for ‘zoonotic’ strains of Giardia among domestic and wild animal, and human hosts.

Emergence of Giardia in Wildlife In mammalian wildlife, Giardia has been reported on every continent and from a wide variety of species, including marsupials, rodents, insectivores, ungulates, marine mammals, felids, canids and ursids (Appelbee et al., 2005; Hamnes et al., 2006; S.J. Kutz et al., unpublished). Increasingly, efforts have been made to determine the strain and zoonotic potential of Giardia, and this has resulted in new host and geographical records for Giardia duodenalis assemblage A in muskoxen (Kutz et al., 2008), bowhead whale (Hughes-Hanks et al., 2005), harp seals (M. Olson, 2007, Calgary, personal communication) and gorillas (Graczyk et al., 2002); assemblage B in beavers (Fayer et al., 2006) and coyotes (Trout et al., 2006); and assemblages A and B from red foxes (Hamnes et al., 2007), ringed seals (B. Dixon and M. Olson, 2007, Morelia, personal communication) and house mice ((Moro et al., 2003). Additional reports of Giardia spp. or Giardia duodenalis in wildlife are common (Hamnes et al., 2006; Gaydos et al., 2007), but unfortunately, without genotyping, the interpretation of these findings is limited with respect to understanding parasite flow and impacts.

Some Case Studies: ‘Zoonotic’ Giardia in Free-ranging Ungulates In general, ungulates typically appear to be accidental hosts for Giardia assemblage A. This is supported by the studies on domestic cattle that invariably report assemblage E as the dominant strain. Giardia is rare in free-ranging ungulate populations (see Hamnes et al., 2006), but zoonotic strains have been recovered from white-tailed deer, elk and muskoxen (Deng and Cliver, 1999; Trout et al., 2003; Hamnes et al., 2006) (S.J. Kutz et al., unpublished). Two studies illustrate some of the key features of the ecology of Giardia that may have significance in the transmission of the parasite to people, and are worthy of further consideration.

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Giardia in Norwegian cervids Giardia is emerging as a parasite of concern in people and animals in Norway (Robertson and Gjerde, 2006). In a study of the parasite in cervids, Giardia was found in red deer (prevalence 1.7%), reindeer (7.1%), moose (12.3%), and roe deer (15.5%) (Hamnes et al., 2006). The difference in prevalence among species was hypothesized to be related to host behaviour and distribution. The higher prevalence in moose was linked to the aquatic habitat that this species prefer and the higher prevalence in roe deer to the fact that the deer are relatively sedentary and commonly found in close proximity to people and agricultural lands (Hamnes et al., 2006). In contrast, reindeer were found in higher, drier and less populated areas, and both red deer and reindeer are migratory species (in contrast to moose and roe deer) and are less likely to re-infect themselves from contaminated soil and water. Thus, this study begins to unravel the potential role of host ecology in the structure of a parasite web for Giardia and in the rates of parasite flow.

Giardia in muskoxen in an arctic ecosystem Giardia was first found in wild muskoxen in 1994 on Banks Island, in arctic Canada. It was found that 4% of muskoxen tested from 1994 to 1998 were positive for the parasite, but the strain was not determined (Nagy et al., 1998). In 2004, muskoxen were sampled again and Giardia assemblage A (probably A1) was reported at a prevalence of 21% (Kutz et al., 2008). These results were interesting for two reasons. First, in this remote region it was anticipated that any Giardia in muskoxen would be either a novel strain or a livestock strain; and secondly, the prevalence of infection appeared to have increased from the mid 1990s to 2004. It was hypothesized that people first introduced Giardia to muskoxen and that the parasite may now be cycling in the muskox population (Kutz et al., 2008), roe deer, moose and reindeer (van der Giessen et al., 2006; Robertson et al., 2007). The time frame for this introduction is unknown. Muskoxen were rare (perhaps absent) on the island in the early 1900s and were unlikely to maintain a population density sufficient to maintain the parasite. Thus the introduction of Giardia may more likely have occurred within the last century. Giardia has also been reported in sympatric Peary caribou (prevalence 3%) as well as in residents of the island. Chronic ‘stomach flu’ has often been reported by people in the community of Sachs Harbour on Banks Island and, since the discovery of Giardia in muskoxen, faecal samples from symptomatic people have been examined for Giardia with some positive results. Ringed seals in the marine environment surrounding the island have also tested positive for Giardia (Olson et al., 1997) and assemblage A has been isolated from some of these animals (M. Olson, 2007, Calgary, personal communication), suggesting a possible terrestrial–marine cycle. The isolates of Giardia from people on Banks Island have not yet been typed and this hinders our understanding of the flow of the parasite among wildlife and people. Several aspects of human and animal behaviour may influence parasite

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transmission in this ecosystem. First, the residents of Banks Island still maintain close ties with the land and spend time ‘on-the-land’ engaged in subsistence hunting and fishing. They visit and camp in areas rich in wildlife, such as river valleys, and obtain water directly from the rivers and streams. The limited summer tourism on the island is also concentrated around waterbodies. Typically there are no latrines in these remote regions and environmental contamination with human waste is common. Additionally, muskoxen on the island are relatively sedentary and are concentrated in the lush river valleys. Thus spatial overlap of habitat use between people and muskoxen is common and is usually centred on waterbodies – ideal locations for the persistence and transmission of Giardia (Kutz et al., 2008). Second, annually during the winter there is an organized, inspected commercial slaughter of muskoxen for sale of meat and qiviut (undercoat). Muskoxen are herded into a central location, held for up to 72 h, and then shot. Historically, the offal from the slaughter was removed to a nearby hillside, but practices have changed and it is now deposited on the sea ice. Depending on survival of the Giardia cysts in the sub-zero environment, the concentration of muskoxen at the slaughter site and the disposal of offal on the land and sea ice may enhance transmission and dispersal of Giardia among the marine and terrestrial wildlife as well as people (Kutz et al., 2008). Preliminary work suggests that the muskox is a good host for Giardia assemblage A, not unlike the beaver. The high prevalence in muskoxen in an area of such low human density is perhaps surprising and suggests that the parasite may be maintained in this host in the absence of people. Possible reduced immunocompetence of muskoxen (this species seems to be exquisitely sensitive to many pathogens) together with enhanced survival and transmission of parasite cysts in an arctic environment may in part explain this pattern of infection. Many questions remain as to how Giardia is maintained in this arctic environment: is it amplified in the muskox population, did it originate from people (spillover) and is there spill-back to people, what other wildlife species are infected, and is there a terrestrial/marine cycle? Giardia may be emerging in muskoxen (increased prevalence since the mid-1990s), but more long-term data on seasonal, temporal, age- and sex-related patterns of infection are needed to confirm this. Further work is planned in order to better understand the flow, effects, patterns of survival and transmission of Giardia in terrestrial and marine environments at these high latitudes (Kutz et al., 2008).

‘Zoonotic’ Giardia in Other Wildlife Hosts Marine ecosystems Reports of Giardia in marine environments are becoming more common. The parasite has been reported in a variety of pinnipeds, and more recently in cetaceans (Olson et al., 1997; Measures and Olson, 1999, 2002; Deng et al., 2000; Hughes-Hanks et al., 2005). Few of these isolates have been typed; however, both assemblage A and B have been recovered from marine mammals

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(Appelbee et al., 2005; B. Dixon, 2007, Morelia, personal communication). In some cases the source of Giardia may be of anthropogenic origin (runoff from human activities) or domestic livestock sources, and the finding of Giardia assemblage A in Macoma and Asian freshwater clams in fresh water and estuaries along the east coast of the USA (Graczyk et al., 1999a, 1999b) supports this hypothesis. Non-human primates Zoonotic transmission of pathogens between human and non-human primates has long been of great concern for human health. With increased ecotourism, the risk of transmission of pathogens from people to non-human primates, and the impact on the conservation of these species, has become a major concern (Nizeyi et al., 2001). Recently, Giardia assemblage A was reported in sympatric gorillas (prevalence of 2%), cattle (10%) and people (5%) in the Bwindi Impenetrable National Park, Uganda. It is hypothesized that the parasite was introduced by people and now circulates among these three host species, and that transmission may be maintained and enhanced by poor husbandry and hygiene practices together with ingestion of untreated stream water (Graczyk et al., 2002).

Drivers of Emergence of Giardia in Wildlife In recent years there has been an apparent increase in the frequency of reports of Giardia in wildlife. This may represent apparent emergence attributable in part to increased sampling efforts related to an interest in the potential role of wildlife as reservoirs for zoonoses, and to improvements in molecular tools for diagnosis. There is, however, strong evidence to suggest that there is a real emergence of human and domestic animal strains of Giardia in wildlife as a result of environmental changes and the breakdown of ecological barriers separating host species. For example, new host assemblages resulting from agricultural encroachment on to wild lands, suburbanization of people and urbanization of wildlife, and pathogen pollution from increased sewage discharge and runoff from agricultural lands, may be linked to a real emergence of Giardia in at least some wildlife species (roe deer and agriculture; gorillas, ecotourism and cattle; and terrestrial runoff to marine systems). Directional climate change, which will continue to alter habitats, habitat use, and water availability and distribution, is likely to result in new host distributions and associations and the subsequent emergence of Giardia strains and other pathogens in new host species and in new locations (Kutz et al., 2004, 2005).

Impacts on Wildlife Although Giardia is recognized as an important pathogen in people and domestic animals, causing gastrointestinal disease and adversely affecting several

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aspects of childhood development (Savioli et al., 2006), effects of Giardia in wildlife have been only superficially investigated. In people and domestic animals Giardia can be treated, and for domestic livestock the impacts of parasitism can often be mitigated through abundant high-quality feed and shelter. For freeranging wildlife, however, this is not the case. Wildlife are often ‘living on the edge’, exposed to the elements, predators, and often with limited food sources. Any additional drain on their resources, such as parasitism, may have significant impacts on the survival and lifetime reproductive success of a free-ranging species.

Host infections with multiple strains It is important to recognize that different strains (historical versus recently acquired) may have different impacts on wildlife hosts. In particular, assemblages A and B may have evolved primarily in a domestic cycle (people/livestock/dogs), and a host switch to wildlife may have more significant impacts on these new hosts. Recent molecular typing has demonstrated that a single host species can be infected with multiple strains of Giardia concurrently. For example, the small Australian marsupial, the quenda, is host to a ‘new’ strain of Giardia which is the only strain present when quenda are in their native habitat (Adams et al., 2004). Quenda in agricultural areas, however, also harbour the livestock strain of Giardia (Thompson, Chapter 1, this volume) indicating host-switching by Giardia and a lack of strong cross-immunity between the two strains. Similarly, coyotes and foxes have recently been reported to harbour several different strains of Giardia in a single host population, and possibly within a single host (Trout et al., 2006; Hamnes et al., 2007). These results raise questions about whether infection with multiple strains of Giardia at population and individual animal levels reflect the historical situation, or whether it is a result of relatively recent increases in contact between previously separated host species. Apparent host switching by Giardia draws attention to the need for an improved understanding of host specificity, the health effects of the parasite in historical and in naïve hosts, how Giardia species and strains are maintained in a variety of ecosystems, and the potential for spillover and spill-back among host species and populations.

Implications for People of ‘Zoonotic Strains’ in Wildlife Although the evidence for wildlife as a significant, ongoing source and reservoir of human infection with Giardia is equivocal, it is conceivable that, once infected, these hosts may serve as a source of ‘spill-back’ of the parasite to people. Beavers and muskoxen, with high reported prevalences and intensities of these strains, and typically an association with waterbodies, stand out as possible wildlife species that may be able to amplify zoonotic Giardia (Fig. 8.1). There may be other, as yet uninvestigated, wildlife hosts with a similar role in the epidemiology of zoonotic Giardia. Despite this potential, there is, at present, no clear evidence that these hosts are a source of human infection, and wildlife may more probably

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Beavers Muskoxen

People

Other wildlife

Domestic animals

Fig. 8.1. A proposed parasite web for Giardia duodenalis assemblages A or B within and among species and between domestic and sylvatic cycles. The thickness of arrows reflects the rate of parasite flow in the various parts of the web, with dashed arrows indicating that parasite flow is unlikely. Note that for beavers and muskoxen a strong ‘within species’ transmission cycle is hypothesized.

be victims of environmental contamination with Giardia from anthropogenic sources as opposed to the primary villains responsible for infections in people. Proposed relative rates of flow between people, domestic animals and wildlife for Giardia assemblages A and B are shown in Fig. 8.1.

Moving Forward Giardia, the ‘cosmopolitan parasite’, is recognized as an important component of the World Health Organization ‘Neglected Disease Initiative’ (Savioli et al., 2006). We propose that, because of the current lack of clarity surrounding the role of wildlife in parasite webs for Giardia, and especially the role of these hosts as sources of infection for people, wildlife should be considered the ‘neglected hosts’ of the ‘Neglected Disease Initiative’. Good datasets on the prevalence, intensity, distribution, and host range of Giardia are rare, and the temporal, spatial and quantitative patterns of parasite flow within and between species (wild, domestic and human) remain poorly described. It is now time to apply our rapidly advancing molecular tools, together with classical epidemiology and parasitology, to better understand the ecology, impacts and flow of Giardia among wild and domestic hosts. This can be done by focusing on several key approaches. Survey and inventory Equipped with molecular techniques to better define species and strain diversity, we now need to revisit host species and geographical regions to (re)define the

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host range and geographical distribution of the various strains of Giardia. Studies that report the presence of ‘Giardia’ with no data on strain identification provide little insight into parasite flow or the epidemiology of the parasite among its various host species. Strain-level identification of Giardia is essential to generate important new knowledge on host–parasite associations and establish baselines for future comparisons. Sampling wildlife poses several logistic challenges, and often there is limited or substandard material available; thus sensitive and efficient diagnostic techniques are essential in order to encourage strain identification (Fayer et al., 2006). Further taxonomic and phylogenetic analyses are also needed to evaluate host–parasite associations and historical biogeographical relationships which should shed light on epidemiology and impacts of the various strains in wildlife and domestic systems.

Classical parasitology Moving beyond basic survey and inventory, there is a need to return to the examples of Davies and Hibler (1979) and others to describe and understand the temporal, spatial, demographic and quantitative patterns of occurrence, survival and transmission of Giardia in wildlife and among wildlife, domestic species and people. Our advantage today is that we can identify strains, greatly enhancing our understanding of observed patterns in wild and experimental settings. Experimental infections of various host species are needed in order to determine the host susceptibility, pre-patent and patent periods, clinical signs and temporal patterns of parasite production. The viability of various Giardia strains from different host species and under different environmental conditions needs to be quantified. Together, these data will provide tremendous insight into the flow of zoonotic strains of Giardia between domestic and wild systems, moving us towards an understanding of causal, as opposed to correlational, relationships.

Giardia archives Finally, there is a need for a concerted effort to archive Giardia isolates from various species and geographical locations. If research in recent years has taught us nothing else, advances in molecular techniques invariably result in the need to go back and re-evaluate previous isolates and species and strain designations. Methodologies and standards for preserving and archiving specimens should be developed and widely promoted to ensure that these valuable materials are available for these retrospective analyses.

Conclusions In the research reported in many peer-reviewed publications, the mere presence of Giardia in wildlife has been sufficient to implicate wildlife as source of infection

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for people. However, the bulk of the evidence to support this claim demonstrates correlation, not causation. In fact, in many cases, wildlife species are more probably the victims, with ‘spillover’ of zoonotic strains from people and/or domestic animals. A critical re-evaluation of zoonotic strains of Giardia in wildlife is needed. This should include a return to classical parasitology and epidemiology to develop a quantitative understanding of the temporal and spatial patterns and impacts of the various Giardia strains in wildlife and the potential interactions with domestic systems. These efforts should lead to an understanding of the structure of parasite webs and parasite flow for Giardia among domestic and sylvatic hosts and people, and should ultimately provide an evidence-based assessment of the critical control points and management activities required to reduce environmental contamination and cross-species transmission. We should not be surprised if we discover that the key management steps needed to reduce Giardia in people may be centred on the appropriate containment and disposal of our own waste!

References Adams, P.J., Monis, P.T., Elliot, A.D. and Thompson, R.C. (2004) Cyst morphology and sequence analysis of the small subunit rDNA and ef1 alpha identifies a novel Giardia genotype in a quenda (Isoodon obesulus) from Western Australia. Infection, Genetics and Evolution 4, 365–370. Appelbee, A., Thorlakson, C. and Olson, M. (2002) Genotypic characterization of Giardia cysts isolated from wild beaver in southern Alberta, Canada. In: Olsen, B.E., Olsen, M.E. and Wallis, P.M. (eds) Giardia: The Cosmopolitan Parasite. CABI Publishing, Wallingford, UK and New York, pp. 299–300. Appelbee, A.J., Thompson, R. and Olson, M.E. (2005) Giardia and Cryptosporidium in mammalian wildlife: current status and future needs. Trends in Parasitology 21, 370–376. Cacciò, S.M., Thompson, R.C.A., McLauchlin, J. and Smith, H.V. (2005) Unravelling Cryptosporidium and Giardia epidemiology. Trends in Parasitology 21, 430–437. Charron, D.F., Thomas, M.K., Waltner-Toews, D., Aramini, J.J., Edge, T., Kent, R.A., Maarouf, A.R. and Wilson, J. (2004) Vulnerability of waterborne diseases to climate change in Canada: a review. Journal of Toxicology and Environmental Health – Part A 67, 1667–1677. Craun, G.F., Calderon, R.L. and Craun, M.F. (2005) Outbreaks associated with recreational water in the United States. International Journal of Environmental Health Research 15, 243–262. Davies, R.B. and Hibler, C.P. (1979) Animal reservoirs and cross-species transmission of Giardia. In: Jakubowski, W. and Hoff, J.C. (eds) Waterborne Transmission of Giardiasis. US Environmental Protection Agency, Office of Research and Development, Environmental Research Centre, Cincinnati, OH, EPA-600/7-79-001, pp. 104–126. Deng, M. and Cliver, D. (1999) Improved immunofluorescence assay for detection of Giardia and Cryptosporidium from asymptomatic adult cervine animals. Parasitology Research 85, 733–736. Deng, M.Q., Peterson, R.P. and Cliver, D.O. (2000) First findings of Cryptosporidium and Giardia in California sea lions (Zalophus californianus). Journal of Parasitology 86, 490–494.

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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. Veterinary Parasitology 135, 105–112. Gaydos, J.K., Miller, W.A., Gilardi, K.V.K., Melli, A., Schwantje, H., Engelstoft, C., Fritz, H. and Conrad, P.A. (2007) Cryptosporidium and Giardia in marine-foraging river otters (Lontra canadensis) from the Puget Sound Georgia Basin ecosystem. Journal of Parasitology 93, 198–202. Graczyk, T.K., Fayer, R., Conn, D.B. and Lewis, E.J. (1999a) Evaluation of the recovery of waterborne Giardia cysts by freshwater clams and cyst detection in clam tissue. Parasitology Research 85, 30–34. Graczyk, T.K., Thompson, R.C., Fayer, R., Adams, P., Morgan, U.M. and Lewis, E.J. (1999b) Giardia duodenalis cysts of genotype A recovered from clams in the Chesapeake Bay subestuary, Rhode River. American Journal of Tropical Medicine and Hygiene 61, 526–529. Graczyk, T.K., Bosco-Nizeyi, J., Ssebide, B., Thompson, R.C., Read, C. and Cranfield, M.R. (2002) Anthropozoonotic Giardia duodenalis genotype (assemblage) A infections in habitats of free-ranging human-habituated gorillas, Uganda. Journal of Parasitology 88, 905–909. 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. Veterinary Parasitology 141, 30–41. Hamnes, I.S., Gjerde, B.K., Forberg, T. and Robertson, L.J. (2007) Occurrence of Giardia and Cryptosporidium in Norwegian red foxes (Vulpes vulpes). Veterinary Parasitology 143, 347–353. Hughes-Hanks, J.M., Rickard, L.G., Panuska, C., Saucier, J.R., O’Hara, T.M., Dehn, L. and Rolland, R.M. (2005) Prevalence of Cryptosporidium spp. and Giardia spp. in five marine mammal species. Journal of Parasitology 91, 1225–1228. Kutz, S.J., Hoberg, E.P., Nagy, J., Polley, L. and Elkin, B. (2004) “Emerging” parasitic infections in arctic ungulates. Integrative and Comparative Biology 44, 109–118. Kutz, S.J., Hoberg, E.P., Polley, L. and Jenkins, E. (2005) Global warming is changing the dynamics of arctic host–parasite systems. Proceedings of the Royal Society B: Biological Sciences 272(1581), 2571–2576. Kutz, S.J., Thompson, R.C.A., Polley, L., Kandola, K., Nagy, J., Wielinga, C.M. and Elkin, B.T. (2008) Giardia assemblage A: human genotype in muskoxen in the Canadian Arctic. Parasites and Vectors. Available at http://www.biomedcentral.com/ (Doi: 10.1186/1756-3305-1-32.) Measures, L.N. and Olson, M. (1999) Giardiasis in pinnipeds from eastern Canada. Journal of Wildlife Diseases 35, 779–782. Measures, L.N. and Olson, M.E. (2002) Giardiasis in Canadian phocid seals. In: Olsen, B.E., Olsen, M.E. and Wallis, P.M. (eds) Giardia: The Cosmopolitan Parasite. CAB International, Wallingford, UK and New York, p. 316. Moro, D., Lawson, M.A., Hobbs, R.P. and Thompson, R.C.A. (2003) Pathogens of house mice on arid Boullanger Island and subantarctic Macquarie Island, Australia. Journal of Wildlife Diseases 39, 762–771. Nagy, J.A., Larter, N., Branigan, M., McLean, E. and Hines, J. (1998) Co-management Plan for Caribou Muskoxen, Arctic Wolves, Snow Geese, and Small Herbivores on Banks Island, Inuvialuit Settlement Region, Northwest Territories. Department of Resources, Wildlife, and Economic Development for the Wildlife Management Advisory Council, Inuvik, Northwest Territories, Canada.

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S.J. Kutz et al. Nizeyi, J.B., Innocent, R.B., Erume, J., Kalema, G.R., Cranfield, M.R. and Graczyk, T.K. (2001) Campylobacteriosis, salmonellosis, and shigellosis in free-ranging humanhabituated mountain gorillas of Uganda. Journal of Wildlife Diseases 37, 239–244. Olson, M.E., Roach, P.D., Stabler, M. and Chan, W. (1997) Giardiasis in ringed seals from the Western Arctic. Journal of Wildlife Diseases 33, 646–648. Parkinson, A.J. and Butler, J.C. (2005) Potential impacts of climate change on infectious diseases in the Arctic. International Journal of Circumpolar Health 64, 478–486. Roach, P.D., Olson, M.E., Whitley, G. and Wallis, P.M. (1993) Waterborne Giardia cysts and Cryptosporidium oocysts in the Yukon, Canada. Applied and Environmental Microbiology 59, 67–73. Robertson, L.J. and Gjerde, B.K. (2006) Fate of Cryptosporidium oocysts and Giardia cysts in the Norwegian aquatic environment over winter. Microbial Ecology 52, 597–602. Robertson, L.J., Forberg, T., Hermansen, L., Hamnes, I.S. and Gjerde, B. (2007) Giardia duodenalis cysts isolated from wild moose and reindeer in Norway: genetic characterization by PCR-rflp and sequence analysis at two genes. Journal of Wildlife Disease 43, 576–585. Savioli, L., Smith, H. and Thompson, A. (2006) Giardia and Cryptosporidium join the ‘Neglected Diseases Initiative’. Trends in Parasitology 22, 203–208. Sulaiman, I.M., Fayer, R., Bern, C., Gilman, R.H., Trout, J.M., Schantz, P.M., Das, P., Lal, A.A. and Xiao, L. (2003) Triosephosphate isomerase gene characterization and potential zoonotic transmission of Giardia duodenalis. Emerging Infectious Diseases 9, 1444–1452. Thompson, R.C.A. (2002) Towards a better understanding of host specificity and the transmission of Giardia: the impact of molecular epidemiology. In: Olsen, B.E., Olsen, M.E. and Wallis, P.M. (eds) Giardia: The Cosmopolitan Parasite. CAB International, Wallingford, UK and New York, pp. 55–69. Trout, J., Santín, M. and Fayer, R. (2003) Identification of assemblage A Giardia in whitetailed deer. Journal of Parasitology 89, 1254–1255. Trout, J.M., Santín, M. and Fayer, R. (2006) Giardia and Cryptosporidium species and genotypes in coyotes (Canis latrans). Journal of Zoo and Wildlife Medicine 37, 141–144. van der Giessen, J.W.B., de Vries, A., Roos, M., Wielinga, P., Korteek, L.M. and Mank, T.G. (2006) Genotyping Giardia in Dutch patients and animals: a phylogenic analysis of human and animal isolates. International Journal for Parasitology 36, 849– 858.

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The Role of Livestock in the Foodborne Transmission of Giardia duodenalis and Cryptosporidium spp. to Humans B.R. DIXON Health Canada, Ottawa, Canada

Abstract While person-to-person and waterborne transmission probably account for most human infections with Giardia duodenalis and Cryptosporidium spp., zoonotic transmission, particularly from livestock, has generated a great deal of interest in recent years. Both G. duodenalis and Cryptosporidium spp. are highly prevalent in cattle and other livestock, and zoonotic genotypes and species of these parasites have been identified in numerous studies. There is limited evidence, however, supporting the role of livestock as reservoirs for human infection. The association of livestock with the foodborne transmission of these parasites is also poorly recognized. Nevertheless, these animals have been associated with the presence of Giardia cysts and Cryptosporidium oocysts in a variety of foods, as well as with some foodborne outbreaks. The possible role of livestock in the contamination of produce, meats and other foods with G. duodenalis and Cryptosporidium spp., and its public health significance will be discussed.

Introduction Giardia and Cryptosporidium are common protozoan parasites of the intestinal tract of humans and a wide variety of animals worldwide. They represent the most common causes of enteric illness associated with protozoan parasites in humans. Giardia and Cryptosporidium are also responsible for illness and production losses in domestic animals (Olson et al., 2003; Olson et al., 2004, O’Handley and Olson, 2006). Transmission of Giardia and Cryptosporidium occurs by means of the faecal–oral route. Giardia cysts and Cryptosporidium oocysts are either ingested directly, as in person-to-person transmission, or indirectly by means of a vehicle such as contaminated water or food. Numerous waterborne outbreaks have been documented worldwide and this has, in fact, been recognized as one of the most common means of transmission for these © CAB International 2009. Giardia and Cryptosporidium: From Molecules to Disease (eds G. Ortega-Pierres et al.)

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pathogens. A number of foodborne illness outbreaks associated with Giardia and Cryptosporidium, and involving a variety of foods, have also been reported. In recent years, there has been considerable interest in the high prevalence of both Giardia and Cryptosporidium infections in livestock, and the possible role these animals may play in the zoonotic transmission of these parasites.

Taxonomy Giardia There are currently six recognized species of Giardia (Olson et al., 2004, Thompson and Monis, 2004; Cacciò et al., 2005). Three of these species, G. duodenalis, G. muris and G. microti, are infectious to mammals, but only G. duodenalis has a broad host range, infecting a wide range of mammals including humans, livestock and companion animals, and is the only species of public health concern. Numerous genotypes, or assemblages, as well as subgroups or subgenotypes within these assemblages, have been demonstrated for G. duodenalis (Thompson, 2004; Thompson and Monis, 2004; Lalle et al., 2005). Thompson (2004) suggested that the genetic distance separating some of these assemblages, including assemblages A and B, is high enough to warrant separating them into different species. Humans are only susceptible to infection with two of these genotypes, assemblages A and B (Cacciò et al., 2005). Cattle and other livestock were thought to be susceptible to infection only with the so-called zoonotic genotype, assemblage A, and with the hoofed livestock genotype, assemblage E (Olson et al., 2004). A recent study, however, has demonstrated that cattle can also be infected with assemblage B (Lalle et al., 2005).

Cryptosporidium The taxonomy of Cryptosporidium is in a constant state of flux. Recently, Fayer (2004) recognized a total of 15 species of Cryptosporidium infecting fish, amphibians, reptiles, birds and mammals. These species are discussed in depth by Ryan (2003) and by Xiao et al. (2004). At least two other species, C. suis (Ryan et al., 2004) and C. bovis (Fayer et al., 2005), have been described since then. There have also been numerous host-specific genotypes of the ‘C. parvum group’ which have been reported in recent years, some of which may represent distinct species (Ryan, 2003). For many years, C. parvum was thought to be the only species infective to humans. However, many additional Cryptosporidium species have recently been identified as causing infection and illness in both immunocompromised and immunocompetent humans (Xiao et al., 2003, 2004; Cacciò et al., 2005). Currently, at least seven species and two genotypes of Cryptosporidium are associated with human disease (Cacciò et al., 2005). The two species of most significance in terms of public health, however, are C. parvum and C. hominis. C. parvum, previously known as the bovine genotype, has a very wide host range, including humans and livestock, and is thought to be the most important

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species in terms of zoonotic transmission. C. hominis is predominantly found in humans; however, there have been some rare natural infections reported in cattle and sheep, as well as some experimental infections (Ryan et al., 2005; Smith et al., 2005).

Transmission Routes The direct faecal–oral route (person-to-person) and the indirect transmission of Giardia cysts and Cryptosporidium oocysts by means of contaminated water probably account for the majority of human cases worldwide. Person-to-person transmission has been identified as the cause of many outbreaks, and is largely associated with poor hygiene. Outbreaks at daycare facilities, in particular, have been the focus of much attention. A large number of surveys have been done on surface and groundwater worldwide, and have demonstrated a relatively high prevalence of both Cryptosporidium and Giardia. As a result, numerous drinking-water-related outbreaks of Cryptosporidium (Fayer, 2004) and Giardia (Jakubowski and Craun, 2002) have also been reported. In recent years, outbreaks, particularly of Cryptosporidium infections, have also been associated with recreational water, including swimming pools, splash pads and water parks (Fayer, 2004; Smith et al., 2006). A few recent review articles have addressed the issue of foodborne transmission of protozoan parasites, including Giardia and Cryptosporidium (Nichols and Smith, 2002; Dixon, 2003; Dawson, 2005). While the foodborne route of transmission is reported much less frequently, and accounts for fewer outbreaks and cases, than the waterborne route or the direct faecal–oral route, it is becoming more important due to factors such as global trade, international travel, and changes in consumer habits, such as the consumption of more raw and undercooked foods (Dixon, 2003). Zoonotic transmission of Giardia and Cryptosporidium remains a controversial issue, and has become the focus of considerable research in recent years. While the zoonotic significance of companion animals and wildlife remains somewhat questionable (Thompson, 2004; Cacciò et al., 2005), livestock, and cattle in particular, have been identified as a potential significant source of environmental contamination with Giardia cysts and Cryptosporidium oocysts, and subsequent human infections. This is largely based on the high infection rates, and the presence of zoonotic genotypes and species of Giardia and Cryptosporidium in these animals. Zoonotic transmission may occur through direct contact with cysts or oocysts in animal faeces, as in the case of animal handlers, veterinarians, farmers and their families, and farm or petting zoo visitors. Zoonotic transmission, however, may also occur by means of a vehicle such as water or food. Surface or groundwater, for example, may become contaminated through the application of fresh manure to fertilize agricultural land or through agricultural runoff (Heitman et al., 2002; Olson et al., 2004). Similarly, produce may become contaminated directly through the use of manure application to crop lands, or indirectly through irrigation or processing with contaminated water. There is also some evidence for the presence of Giardia cysts and Cryptosporidium oocysts in products of animal origin, such as milk and meat.

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Prevalence of Giardia and Cryptosporidium in Livestock A high prevalence of both Giardia and Cryptosporidium has been reported worldwide in both dairy and beef cattle (Olson et al., 2003). O’Handley (2002), however, indicated that prevalence rates can vary among different studies and that this may result from differences in climate, farm management, study design and diagnostic test used. A large number of prevalence and molecular characterization studies have been done worldwide in recent years on Giardia and Cryptosporidium infections in dairy calves. The prevalence of both parasites has generally been reported to be high, with some studies reporting a 100% cumulative prevalence of both Giardia and Cryptosporidium in dairy calves (Xiao and Herd, 1994; O’Handley et al., 1999; O’Handley, 2002). Studies in Canada have reported a prevalence in dairy calves of between 45.7% and 73% for G. duodenalis, and between 40.6% and 88.7% for Cryptosporidium spp. (Olson et al., 1997b; Ruest et al., 1998; O’Handley et al., 2000; Trotz-Williams et al., 2005). In the USA, Cryptosporidium spp. prevalences of between 7.5% and 49% have been reported in dairy calves (Garber et al., 1994; Nydam et al., 2005; Starkey et al., 2005). Trout et al. (2004, 2005) examined the prevalence and genotypes of G. duodenalis in pre- and post-weaned dairy calves from several states in the USA. These authors reported a widely varying prevalence of 9–93%, with an average of 40% in preweaned calves and an overall prevalence of 52% in post-weaned calves. Numerous prevalence studies in dairy calves have also been undertaken in Europe, primarily on Cryptosporidium, and have demonstrated a Cryptosporidium prevalence ranging from 2.4% to 47.8%, while Giardia prevalence ranged from 0.8% to 49% (Lefay et al., 2000; Huetink et al., 2001; Castro-Hermida et al., 2002; Joachim et al., 2003; Hamnes et al., 2006; Thompson et al., 2007). In India, Singh et al. (2006) reported a Cryptosporidium prevalence in dairy calves of 50% in diarrhoeic calves and 25.7% in non-diarrhoeic calves. Becher et al. (2004) reported Cryptosporidium and Giardia in 48% and 89% of sampled dairy calves, respectively, in Western Australia. Several studies have examined Cryptosporidium and Giardia infections in older dairy cattle, and have reported a generally lower prevalence than in calves (Fayer et al., 2000; Santín et al., 2004; Fayer et al., 2006, 2007). Conversely, Trout et al. (2006) reported a relatively high and varied prevalence of Giardia in 1–2-year-old dairy cattle. Uehlinger et al. (2006) also reported a high prevalence of Giardia in adult dairy cattle in a veterinary college teaching herd, along with a complete absence of Cryptosporidium spp. There have been generally fewer studies on the prevalence of Giardia and Cryptosporidium in beef cattle. Ralston et al. (2003) reported a Cryptosporidium prevalence of only 5% in beef calves in Alberta, Canada. A low prevalence of Cryptosporidium in beef cows and calves was also reported in western Canada by Gow and Waldner (2006). C. parvum has been reported to be considerably less prevalent in beef calves than in dairy calves (Kvac et al., 2006; O’Handley and Olson, 2006). Conversely, some studies have demonstrated a relatively high Cryptosporidium prevalence in beef cattle (Fayer et al., 2000; McAllister et al., 2005). For Giardia, Ralston et al. (2002, 2003) reported a 100% cumulative

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prevalence in beef calves in Alberta, Canada, while more moderate Giardia infection rates have been reported for beef cattle in other studies (Fayer et al., 2000; Appelbee et al., 2003; McAllister et al., 2005; Gow and Waldner, 2006). A relatively high prevalence of both Giardia and Cryptosporidium has also been reported worldwide in swine (Olson et al., 1997a; Ryan et al., 2003; Xiao et al., 2006). As with dairy and beef calves, a 100% cumulative prevalence of Cryptosporidium in pigs has also been reported (Guselle et al., 2003). A number of studies have been done worldwide on the prevalence of Giardia and Cryptosporidium in sheep and have reported prevalences of between 6.2% and 68.6% for Giardia, and between 10.1% and 68.3% for Cryptosporidium (Olson et al., 1997a; Ryan et al., 2005). Santín et al. (2007) reported a much higher prevalence of Cryptosporidium in lambs (77.4%) than in ewes (25%) on a sheep farm in Maryland, USA, and a relatively low prevalence of Giardia in both lambs (4%) and ewes (12%). A few recent studies have also reported the presence of Giardia and Cryptosporidium in other livestock including horses (Olson et al., 1997a; Atwill et al., 2000), and goats (Bomfim et al., 2005; Castro-Hermida et al., 2005).

Zoonotic Genotypes and Species of Giardia and Cryptosporidium in Livestock Giardia duodenalis Cattle may be infected with either the zoonotic G. duodenalis assemblage A or the host-adapted, non-zoonotic, assemblage E. Numerous studies have reported a higher prevalence of G. duodenalis assemblage E than assemblage A in cattle (O’Handley et al., 2000; Appelbee et al., 2003; Trout et al., 2004, 2005, 2006; O’Handley and Olson, 2006), and Olson et al. (2004) concluded that this probably limits their role as reservoirs for human infection. However, a similar prevalence of both assemblages was reported by Uehlinger et al. (2006) in adult dairy cattle in a veterinary college teaching herd, and is suggestive of a potential zoonotic risk of infection to humans. Common genotypes or species in different hosts is not necessarily evidence for zoonotic transmission, however (Thompson, 2002). O’Handley (2002) suggested that in order to understand the importance of livestock in the zoonotic transmission of Giardia, it must be determined whether the genotypes infectious to humans are naturally transmitted among livestock, or whether livestock become infected with these genotypes through contact with a human or other animal source. Recently, O’Handley and Olson (2006) indicated that the presence of assemblage A in ruminants may, in fact, be suggestive of a human source of infection. It seems feasible, therefore, that the relatively high proportion of assemblage A in cattle from a teaching herd reported by Uehlinger et al. (2006) may be evidence for human-to-animal transmission, particularly given the frequent human contact with these animals. Recently, cattle have also been reported to be infected with G. duodenalis assemblage B (Lalle et al., 2005), greatly increasing the potential reservoir of infection for humans, and raising questions about the actual source of infection

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in these animals. Recent unpublished results from our laboratory on the molecular characterization of Giardia in livestock support these findings and have, in fact, demonstrated a predominance of assemblage B in pooled dairy cattle manure and in pooled swine manure in eastern Ontario, Canada. Specifically, we found a predominance of G. duodenalis assemblage B in pooled swine manure, with the remainder being assemblage E. In dairy cattle, we identified both G. duodenalis assemblages B and E, as well as a small number of samples which typed as assemblage A. This predominance of assemblage B is very suggestive of humanto-animal transmission, but it is not clear whether this genotype may, in fact, be naturally transmitted among livestock in this region. Traub et al. (2005) examined the genotypes of G. duodenalis in horses from New York and from Western Australia, and reported the presence of assemblage AI in Australia and AI, AII and BIV in New York. The authors concluded that horses may constitute a risk of zoonotic transmission either through direct contact with oocysts or via watersheds.

Cryptosporidium spp. Cattle are susceptible to infection with at least four species of Cryptosporidium (Xiao et al. (2007). These include the zoonotic species, C. parvum, as well as the non-zoonotic host-specific species and genotypes C. andersoni, Cryptosporidium deer-like genotype and the recently described C. bovis (Fayer et al., 2005). Santín et al. (2004) demonstrated that infections with these species in cattle are age-related, with C. parvum predominating in pre-weaned calves, and being replaced with other Cryptosporidium species, including C. andersoni, C. bovis and the deer-like genotype, in post-weaned calves and older animals. Similar findings have been reported by Fayer et al. (2006) and Thompson et al. (2007). Recently, however, Feng et al. (2007) reported that both C. bovis and the Cryptosporidium deer-like genotype were found in all age groups of cattle in different parts of the world. Based on the high prevalence of C. parvum in young animals, Xiao et al. (2007) concluded that only neonatal calves are an important source of zoonotic transmission. Similarly, due to the very low prevalence of C. parvum in mature dairy cattle, Fayer et al. (2007) suggested that these animals pose a low risk of infection to humans. In pigs, two distinct Cryptosporidium genotypes have been reported, pig genotype I and pig genotype II (Ryan et al., 2003). Ryan et al. (2004) determined that pig genotype I actually constituted a new species, and named it C. suis. Cryptosporidium suis has subsequently been included in the list of Cryptosporidium species infective to humans. Guselle et al. (2003) reported the presence of C. suis in all pigs tested in Alberta, Canada. Xiao et al. (2006) identified C. suis, Cryptosporidium pig genotype II, and another potentially zoonotic species, C. muris, in pig slurry samples in Ireland, and concluded that spreading pig slurry onto agricultural land could result in contamination of water sources with zoonotic isolates of Cryptosporidium. Recent unpublished results from our laboratory demonstrated a predominance of C. parvum and Cryptosporidium pig genotype II in pooled swine manure, while only one isolate was identified as C. suis.

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Ryan et al. (2005) noted that, with the exception of one C. hominis isolate, most sheep tested in Australia were infected with species and genotypes rarely or never reported in humans, and these authors concluded that sheep may not represent an important zoonotic reservoir for Giardia and Cryptosporidium infections in humans. More recently, Santín et al. (2007) found that species and genotypes in sheep included mainly a novel C. bovis-like genotype, and Cryptosporidium cervine genotype, with only two lambs exhibiting C. parvum. These authors further reported that most Giardia infections were assemblage E, with only one ewe having an infection with assemblage A. They concluded that although there was a low prevalence of C. parvum and G. duodenalis assemblage A, the high prevalence of the zoonotic Cryptosporidium cervine genotype indicated that sheep may play a role in transmission to humans through environmental contamination.

The Significance of Zoonotic Transmission from Livestock While a high prevalence of both Giardia and Cryptosporidium has been reported worldwide in cattle and other livestock, and common genotypes and species exist in humans and livestock, direct evidence for zoonotic transmission remains rather scant (Olson et al., 2004; Thompson, 2004). For example, a waterborne outbreak of cryptosporidiosis in British Columbia, Canada in 1996 is the only waterborne outbreak in North America in which oocysts of the bovine genotype have been identified (Cacciò et al., 2005). Waterborne outbreaks associated with livestock are, in fact, largely based on epidemiological or circumstantial evidence (Olson et al., 2004, Thompson, 2004). Based on the absence of C. hominis infection in dairy calves from farms near New York City, USA, Nydam et al. (2005) concluded that cattle are probably not an important source of infection in waterborne outbreaks, in which C. hominis is generally identified in North America. However, in a recent study, Feltus et al. (2006) identified C. parvum as the predominant cause of sporadic human cases in Wisconsin, USA, and suggested that this was indicative of zoonotic transmission, with cattle and other ruminants being the likely sources based on subtyping data. Further evidence for zoonotic transmission include a foodborne outbreak in which Cryptosporidium-infected calves were thought to be directly associated with oocyst-contaminated apple cider in Maine, USA (Millard et al., 1994). Outbreaks of infection with the bovine genotype (C. parvum) have occasionally been epidemiologically linked to direct or indirect contact with animals (Ryan, 2003, Olson et al., 2004). These included an outbreak among a rural family in Pennsylvania, USA, in 1997, and an outbreak involving a water fountain at the Minnesota Zoo in 1997. Despite this evidence in support of zoonotic transmission, recent studies involving molecular epidemiology have, in fact, suggested that the role of cattle and other livestock in human infections with Giardia and Cryptosporidium may not be as important as was originally thought (O’Handley, 2002; Olson et al., 2004; O’Handley and Olson, 2006). O’Handley and Olson (2006) suggested that only young ruminants pose a risk for zoonotic transmission of Cryptosporidium, and that, overall, human-to-human transmission may be more important than zoonotic transmission.

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The Role of Livestock in Foodborne Transmission of Giardia and Cryptosporidium While a number of different foods have been epidemiologically associated with outbreaks of infection with Giardia and Cryptosporidium, including fresh produce, apple cider, salads and other foods (Dixon, 2003), the actual source of contamination of the food is often not clear. Contamination by infected farm workers, irrigation and/or wash water containing cysts or oocysts, or food handlers, has been suspected in some outbreaks. Another potential source of contamination of food, however, is by means of direct or indirect contact with livestock. Laberge and Griffiths (1996) suggested that the high prevalence of Cryptosporidium spp. in domestic animals directly or indirectly increases the risk for contamination of foods including produce, milk and meat. Foodborne outbreaks associated with livestock are rarely reported but warrant further investigation as to how the foods actually become contaminated, and what can be done to reduce the risk of contamination. Poor farm management practices, for example, may allow livestock access to crops (direct contamination) or surface water (indirect contamination). The irrigation of crops, or the washing or processing of produce, with Giardia- or Cryptosporidium-contaminated raw water represents a significant risk for foodborne transmission. The application of manure onto crop lands is another important source of contamination of fresh produce (Olson et al., 2004; Moore et al., 2007). Giardia cysts and Cryptosporidium oocysts are environmentally resistant and can remain viable in water, soil or manure for extended periods (Olson et al., 1999). Giardia cysts and, particularly, Cryptosporidium oocysts are also very resistant to the environmental conditions found in the field, as well as those during storage or transportation of produce (Dixon, 2003). Further, the moist surface of some fruits and vegetables may, in fact, further protect any contaminating cysts or oocysts from desiccation and unfavourable temperatures. As such, it cannot be assumed that cysts and oocysts contaminating produce in the field, or during storage, will become inactivated before reaching the consumer. In light of the high prevalence of both Giardia and Cryptosporidium in livestock, it is also feasible that carcasses or cuts of meat may become contaminated in the slaughterhouse, either directly by faecal contact or by means of washing or processing with contaminated water.

Prevalence of Giardia and Cryptosporidium in Foods Surveys have reported a relatively high prevalence of Cryptosporidium oocysts on market produce in Costa Rica and Peru (Monge and Chinchilla, 1996; Monge et al., 1996; Ortega et al., 1997; Calvo et al., 2004). Ortega et al. (1997) reported that 14.5% of the total vegetables examined from markets in Peru contained oocysts, and suggested that the consumption of raw or undercooked produce purchased at these markets could be a source of transmission to humans. Giardia cysts have been detected in a variety of foods, including Christmas pudding, lettuce and strawberries (Barnard and Jackson, 1984; Boreham, 1987;

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Boreham et al., 1990). Giardia cysts were reported in 1% of vegetables obtained from supermarkets and a public market in the Philippines (de Leon et al., 1992). The authors speculated that the vegetables may have been contaminated through the application of human and animal manure on agricultural lands, among other sources. Giardia cysts were also reported in market vegetables tested in Costa Rica (Monge et al., 1996). A recent survey done in Norway showed a relatively high prevalence of both Cryptosporidium oocysts and Giardia cysts in a variety of fruits and vegetables, as well as in water samples related to irrigation and production (Robertson and Gjerde, 2001). There is very little information regarding the prevalence of Giardia or Cryptosporidium in or on raw meats. Moriarty et al. (2005) investigated the prevalence of Cryptosporidium in the faeces and on the carcasses of individual beef cattle at slaughter. While Cryptosporidium spp. oocysts were detected in 7.3% of the faecal samples, no Cryptosporidium oocysts were detected in any of the carcass tissue. Similarly, Cryptosporidium oocysts have been detected in adult cattle in slaughterhouses in Japan (Kaneta and Nakai, 1998; Koyama et al., 2005). Recent unpublished results from our laboratory demonstrated the presence of both G. duodenalis and Cryptosporidium spp. on raw meats purchased from retail outlets. Chicken breasts, minced beef and pork chops were all positive for both parasites by nested PCR, while one minced beef sample was positive for Cryptosporidium by immunofluorescence microscopy and one pork chop sample was positive for Giardia using this method. All Giardia isolates in meats were determined by DNA sequence alignment to be G. duodenalis assemblage B, with the exception of one minced beef isolate which belonged to assemblage E. The identification of assemblage B in meats is indicative of a human source of contamination rather than an animal source. In addition, C. parvum was identified in all meats by PCR sequencing positive samples. These results represent the first report of Giardia and Cryptosporidium on naturally contaminated raw meats. A number of recent studies have also reported on the presence and survival of Cryptosporidium oocysts in tissues of oysters and other molluscan shellfish, and the potential for transmission to humans through consumption (Graczyk, 2003; Fayer et al., 2004). Giardia cysts have also been isolated from clams in a sub-estuary of Chesapeake Bay, USA (Graczyk et al., 1999). Along with sewage discharge, agricultural runoff probably contributes significantly to the levels of cysts and oocysts detected in seawater and subsequently in shellfish.

Foodborne Outbreaks Associated with Livestock Approximately 10% of all cases of cryptosporidiosis in the USA have been attributed to foodborne transmission (Mead et al., 1999). Many of these outbreaks have been associated directly or indirectly with livestock. For example, earlier foodborne outbreaks of cryptosporidiosis reported in the UK and Australia, and in travellers to Mexico, were associated with salad, raw milk (cow and goat), raw sausages and raw tripe prepared from the stomach of a cow (Laberge and Griffiths, 1996). More recently, a Cryptosporidium outbreak in Maine, USA, in 1993 was epidemiologically associated with drinking unpasteurized, fresh-pressed apple

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cider (Millard et al., 1994). Oocysts were detected in leftover cider and on swabs from the cider press. It is worth noting that the apples used in making the cider were probably contaminated after falling to the ground near the edge of a pasture from which Cryptosporidium-infected calves were identified. An outbreak of cryptosporidiosis in 1995, involving 50 cases, was reported in the UK and was linked to drinking school milk which had been inadequately pasteurized (Gelletlie et al., 1997). These authors suggested that the milk may have become contaminated through poor udder hygiene. Other recent outbreaks of cryptosporidiosis have been associated with foods such as chicken salad, raw produce and apple cider, but were considered to have been the result of infected food handlers, contaminated water or other unknown sources (Besser-Wiek et al., 1996; Mshar et al., 1997; Quinn et al., 1998; Quiroz et al., 2000). As with cryptosporidiosis, an estimated 10% of all cases of giardiasis in the USA are attributable to foodborne transmission (Mead et al., 1999). There have been numerous foodborne outbreaks of giardiasis reported in North America (Dixon, 2003). The majority of these outbreaks were attributed to foods contaminated by food handlers who are themselves infected, or those who have been in close contact with other infected individuals, particularly young children in diapers. One example of an outbreak of giardiasis with a possible animal origin occurred in Turkey, in which a small number of people became ill following the consumption of tripe soup (Karabiber and Aktas, 1991). The tripe was prepared from sheep, and the authors suggested that the animal could have been infected and that cysts could have remained viable in the crevices of the intestine during cooking.

Conclusions Both Giardia duodenalis and Cryptosporidium spp. are highly prevalent in cattle and other livestock, especially in young animals. Further, molecular characterization has demonstrated that zoonotic genotypes and species of these parasites are common in these animals. However, while zoonotic transmission from livestock to humans has generated considerable interest among researchers in recent years, there remains limited evidence for this form of transmission. Zoonotic transmission by means of the direct or indirect contamination of foods by livestock is less well recognized than other forms, including livestock-contaminated surface water, or direct animal contact. Nevertheless, livestock have been associated with foodborne outbreaks of giardiasis and cryptosporidiosis, and with the presence of cysts and oocysts in a variety of foods. Further work is required to further clarify the role of livestock in the contamination of foods with zoonotic isolates of Giardia and Cryptosporidium, and the control measures that can be taken to minimize this risk.

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Laberge, I. and Griffiths, M.W. (1996) Prevalence, detection and control of Cryptosporidium parvum in food. International Journal of Food Microbiology 31, 1–26. Lalle, M., Pozio, E., Capelli, G., Bruschi, F., Crotti, D. and Cacciò, S.M. (2005) Genetic heterogeneity at the b-giardin locus among human and animal isolates of Giardia duodenalis and identification of potentially zoonotic subgenotypes. International Journal for Parasitology 35, 207–213. Lefay, D., Naciri, M., Poirier, P. and Chermette, R. (2000) Prevalence of Cryptosporidium infection in calves in France. Veterinary Parasitology 89, 1–9. McAllister, T.A., Olson, M.E., Fletch, A., Wetzstein, M. and Entz, T. (2005) Prevalence of Giardia and Cryptosporidium in beef cows in southern Ontario and in beef calves in southern British Columbia. Canadian Veterinary Journal 46, 47–55. 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. Emerging Infectious Diseases 5, 607–625. 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. Journal of the American Medical Association 272, 1592–1596. Monge, R. and Chinchilla, M. (1996) Presence of Cryptosporidium oocysts in fresh vegetables. Journal of Food Protection 59, 202–203. Monge, R., Chinchilla, M. and Reyes, L. (1996) Occurrence of parasites and intestinal bacteria in vegetables that are consumed raw in Costa Rica. Revista de Biologia Tropical 44, 369–375. Moore, J.E., Millar, B.C., Kenny, F., Lowery, C.J., Xiao, L., Rao, J.R., Nicholson, V., Watabe, M., Heaney, N., Sunnotel, O., McCorry, K., Rooney, P.J., Snelling, W.J. and Dooley, J.S.G. (2007) Detection of Cryptosporidium parvum in lettuce. International Journal of Food Science and Technology 42, 385–393. Moriarty, E.M., McEvoy, J.M., Lowery, C.J., Thompson, H.P., Finn, M., Sheridan, J.J., Blair, I.S., McDowell, D.A. and Duffy, G. (2005) Prevalence and characterization of Cryptosporidium species in cattle faeces and on beef carcases at slaughter. Veterinary Record 156, 165–168. Mshar, P.A., Dembek, Z.F., Cartter, M.L., Hadler, J.L., Fiorentino, T.R., Marcus, R.A., Mcguire, J., Shiffrin, M.A., Lewis, A., Feuss, J., Vandyke, J., Toly, M., Cambridge, M., Guzewich, J., Keithly, J., Dziewulski, D., Braunhowland, E., Ackman, D., Smith, P., Coates, J. and Ferrara, J. (1997) Outbreaks of Escherichia coli O157H7 infection and cryptosporidiosis associated with drinking unpasteurized apple cider: Connecticut and New York, October 1996. Journal of the American Medical Association 277, 781–782. Nichols, R.A.B. and Smith, H.V. (2002) Parasites: Cryptosporidium, Giardia and Cyclospora as foodborne pathogens, In: Blackburn, C. de W. and McClure, P.J. (eds) Foodborne Pathogens: Hazards, Risk Analysis and Control. Part III: Non-bacterial and Emerging Foodborne Pathogens. Woodhead Publishing, Cambridge, UK, pp. 453–478. Nydam, D.V., Lindergard, G., Santucci, F., Schaaf, S.L., Wade, S.E. and Mohammed, H.O. (2005) Risk of infection with Cryptosporidium parvum and Cryptosporidium hominis in dairy cattle in the New York City watershed. American Journal of Veterinary Research 66, 413–417. O’Handley, R.M. (2002) Giardia in farm animals. In: Olsen, B.E., Olsen, M.E. and Wallis, P.M. (eds) Giardia: The Cosmopolitan Parasite. CAB International, Wallingford, UK, pp. 97–105. O’Handley, R.M. and Olson, M.E. (2006) Giardiasis and cryptosporidiosis in ruminants. Veterinary Clinics of North America: Food Animal Practice 22, 623–643. O’Handley, R.M., Cockwill, C., McAllister, T.A., Jelinski, M., Morck, D.W. and Olson, M.E. (1999) Duration of naturally acquired giardiosis and cryptosporidiosis in dairy calves

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B.R. Dixon and their association with diarrhea. Journal of the American Veterinary Medical Association 214, 391–396. O’Handley, R.M., Olson, M.E., Fraser, D., Adams, P. and Thompson, R.C. (2000) Prevalence and genotypic characterization of Giardia in dairy calves from Western Australia and western Canada. Veterinary Parasitology 90, 193–200. Olson, M.E., Thorlakson, C.L., Deselliers, L., Morck, D.W. and McAllister, T.A. (1997a) Giardia and Cryptosporidium in Canadian farm animals. Veterinary Parasitology 68, 375–381. Olson, M.E., Guselle, N.J., O’Handley, R.M., Swift, M.L., McAllister, T.A., Jelinski, M.D. and Morck, D.W. (1997b) Giardia and Cryptosporidium in dairy calves in British Columbia. Canadian Veterinary Journal 38, 703–706. Olson, M.E., Goh, J., Phillips, M., Guselle, N. and McAllister, T.A. (1999) Giardia cyst and Cryptosporidium oocyst survival in water, soil, and cattle faeces. Journal of Environmental Quality 28, 1991–1996. Olson, M.E., Ralston, B.J., O’Handley, R., Guselle, N.J. and Appelbee, A.J. (2003) What is the clinical and zoonotic significance of cryptosporidiosis in domestic animals and wildlife. In: Thompson, R.C.A., Armson, A. and Ryan, U.M. (eds) Cryptosporidium: From Molecules to Disease. Elsevier, New York, pp. 51–68. Olson, M.E., O’Handley, R.M., Ralston, B.J., McAllister, T.A. and Thompson, R.C.A. (2004) Update on Cryptosporidium and Giardia infections in cattle. Trends in Parasitology 20, 185–191. 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. American Journal of Tropical Medicine and Hygiene 57, 683–686. Quinn, K., Baldwin, G., Stepak, P., Thorburn, K., Bartleson, C., Goldoft, M., Kobayashi, J. and Stehr-Green, P. (1998) Foodborne outbreak of cryptosporidiosis: Spokane, Washington, 1997. Morbidity and Mortality Weekly Report 47, 565–567. 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. Journal of Infectious Diseases 181, 695–700. Ralston, B.J., McAllister, T.A. and Olson, M.E. (2002) Prevalence and infection pattern of naturally acquired giardiasis in beef calves and their dams from birth to weaning. In: Olsen, B.E., Olsen, M.E. and Wallis, P.M. (eds) Giardia: The Cosmopolitan Parasite. CAB International, Wallingford, UK, pp. 47–52. Ralston, B.J., McAllister, T.A. and Olson, M.E. (2003) Prevalence and infection pattern of naturally acquired giardiasis and cryptosporidiosis in range beef calves and their dams. Veterinary Parasitology 113, 113–122. Robertson, L.J. and Gjerde, B. (2001) Occurrence of parasites on fruits and vegetables in Norway. Journal of Food Protection 64, 1793–1798. Ruest, N., Faubert, G.M. and Couture, Y. (1998) Prevalence and geographical distribution of Giardia spp. and Cryptosporidium spp. in dairy farms in Quebec. Canadian Veterinary Journal 39, 697–700. Ryan, U.M. (2003) Molecular characterization and taxonomy of Cryptosporidium. In: Thompson, R.C.A., Armson, A. and Ryan, U.M. (eds) Cryptosporidium: From Molecules to Disease. Elsevier, New York, pp. 147–160. Ryan, U.M., Samarasinghe, B., Read, C., Buddle, J.R., Robertson, I.D. and Thompson, R.C.A. (2003) Identification of a novel Cryptosporidium genotype in pigs. Applied and Environmental Microbiology 69, 3970–3974. 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

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suis n. sp. (Apicomplexa: Cryptosporidiidae) in pigs (Sus scrofa). Journal of Parasitology 90, 769–773. 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. Applied and Environmental Microbiology 71, 4992–4997. 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. Veterinary Parasitology 122, 103–117. Santín, M., Trout, J.M. and Fayer, R. (2007) Prevalence and molecular characterization of Cryptosporidium and Giardia species and genotypes in sheep in Maryland. Veterinary Parasitology 146, 17–24. Singh, B.B., Sharma, R., Kumar, H., Banga, H.S., Aulakh, R.S., Gill, J.P. and Sharma, J.K. (2006) Prevalence of Cryptosporidium parvum infection in Punjab (India) and its association with diarrhea in neonatal dairy calves. Veterinary Parasitology 140, 162–165. 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. Veterinary Record 156, 710–711. Smith, H.V., Cacciò, S.M., Tait, A., McLauchlin, J. and Thompson, R.C.A. (2006) Tools for investigating the environmental transmission of Cryptosporidium and Giardia infections in humans. Trends in Parasitology 22, 160–167. Starkey, S.R., Wade, S.E., Schaaf, S. and Mohammed, H.O. (2005) Incidence of Cryptosporidium parvum in the dairy cattle population in a New York City watershed. Veterinary Parasitology 131, 197–205. Thompson, H.P., Dooley, J.S.G., Kenny, J., McCoy, M., Lowery, C.J., Moore, J.E. and Xiao, L. (2007) Genotypes and subtypes of Cryptosporidium in neonatal calves in Northern Ireland. Parasitology Research 100, 619–624. Thompson, R.C.A. (2002) Towards a better understanding of host specificity and the transmission of Giardia: the impact of molecular epidemiology. In: Olsen, B.E., Olsen, M.E. and Wallis, P.M. (eds) Giardia: The Cosmopolitan Parasite. CAB International, Wallingford, UK, pp. 55–69. Thompson, R.C.A. (2004) The zoonotic significance and molecular epidemiology of Giardia and giardiasis. Veterinary Parasitology 126, 15–35. Thompson, R.C.A. and Monis, P.T. (2004) Variation in Giardia: implications for taxonomy and epidemiology. Advances in Parasitology 58, 69–137. Traub, R., Wade, S., Read, C., Thompson, A. and Mohammed, H. (2005) Molecular characterization of potentially zoonotic isolates of Giardia duodenalis in horses. Veterinary Parasitology 130, 317–321. Trotz-Williams, L.A., Jarvie, B.D., Martin, S.W., Leslie, K.E. and Peregrine, A.S. (2005) Prevalence of Cryptosporidium parvum infection in southwestern Ontario and its association with diarrhea in neonatal dairy calves. Canadian Veterinary Journal 46, 349–351. Trout, J.M., Santín, M., Greiner, E. and Fayer, R. (2004) Prevalence of Giardia duodenalis genotypes in pre-weaned dairy calves. Veterinary Parasitology 124, 179–186. Trout, J.M., Santín, M., Greiner, E. and Fayer, R. (2005) Prevalence and genotypes of Giardia duodenalis in post-weaned dairy calves. Veterinary Parasitology 130, 177–183. Trout, J.M., Santín, M., Greiner, E.C. and Fayer, R. (2006) Prevalence and genotypes of Giardia duodenalis in 1–2 year old dairy cattle. Veterinary Parasitology 140, 217–222. Uehlinger, F.D., Barkema, H.W., Dixon, B.R., Coklin, T. and O’Handley, R.M. (2006) Giardia duodenalis and Cryptosporidium spp. in a veterinary college bovine teaching herd. Veterinary Parasitology 142, 231–237.

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10

The Risk of Zoonotic Genotypes of Cryptosporidium spp. in Watersheds

H.O. MOHAMMED AND S.E. WADE College of Veterinary Medicine, Cornell University, USA

Abstract Cryptosporidium parvum is a coccidian protozoan that has zoonotic significance. Genotypes of this protozoan are known to contribute significantly to calf morbidity and mortality and hence have become an economic liability for many dairy and beef herds. In addition to its negative impact on the efficiency of animal production, Cryptosporidium has emerged as one of the most significant waterborne pathogens causing enterocolitis in humans worldwide. Because of the broad host spectrum of this pathogen, inefficiency of the common drinking water treatment methods, and the lack of reliable therapy in humans, zoonotic genotypes of Cryptosporidium spp. are considered to be among the major threats found in water supply systems. We have been carrying out studies to evaluate the risk associated with environmental degradation, and hence the contamination of drinking water supply systems with zoonotic Cryptosporidium spp. in a watershed in New York state. There are many agricultural sources that could contribute to the contamination of water supply systems including dairy cattle operations and associated farming activities, wildlife, and sewage treatment plants. The risk assessment approach, based on data collected from this watershed ecosystem, was employed to evaluate the potential environmental degradation. A fault-tree pathway scenario approach was used to assess the likelihood of contamination of the water supply in the watershed. Estimates of the parameters used in the risk assessment model were obtained from our studies and from the literature. The analysis demonstrated the importance of farming activities in mitigating the risk of this protozoan associated with dairy cattle, and highlighted the role of sewage treatment plants.

Introduction Cryptosporidium is a coccidian that can infect most mammals, including humans. Infection is contacted by ingestion of viable oocysts that are shed in the faeces of infected hosts (Fayer et al., 1997). Cryptosporidium has been found to be prevalent in mammals (Atwill et al., 1997; Olson et al., 1997; O’Handley and Olson, © CAB International 2009. Giardia and Cryptosporidium: From Molecules to Disease (eds G. Ortega-Pierres et al.)

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2006; Shukla et al., 2006; Ziegler et al., 2007) including cattle (Garber et al., 1994; Olson et al., 1997; Mohammed et al., 1999; Wade et al., 2000). Most of the dairy herds and wildlife that have been investigated in these studies are inhabitants of the watershed ecosystem. By virtue of their residency in the watershed ecosystem and the likelihood of harbouring genotypes of Cryptosporidium, cattle are likely to pose a risk to human health. This risk can be inferred from the fact that some of these zoonotic genotypes are likely to enter the water supply systems that collect water from the respective watershed and, hence, infect humans. There are several genotypes of Cryptosporidium, some of which are known to be zoonotic. It is plausible that environmental and host factors may have modified the host–parasite ecology and led to the emergence of new pathogenic strains. As a consequence, the parasites have adapted to novel hosts, favouring the survival of ecological generalists (affecting multiple hosts) rather than specialists (affecting one host). At present, 15 distinct species are recognized (Fayer et al., 2005; Hunter and Thompson, 2005). The vast majority of outbreaks of cryptosporidiosis in humans are attributed either to the host-adapted C. hominis or the less fastidious C. parvum (Xiao and Ryan, 2004). C. parvum is known to be prevalent among dairy cattle populations and at a higher rate in calves (Nydam et al., 2001; Santín et al., 2004; Starkey et al., 2006). Cattle also harbour non-zoonotic strains of Cryptosporidium, e.g. C. bovis (Fayer et al., 2005). This organism is microscopically indistinguishable from C. parvum and, therefore, studies which investigated the public health risk associated with dairy animals in watersheds might have overestimated the contributory role of agricultural animals. The dynamics of Cryptosporidium infection in wildlife is not fully understood (Appelbee et al., 2005). Recent studies have incriminated some wildlife species as potential reservoirs of zoonotic genotypes of Cryptospordium (Daszak et al., 2000; Kruse et al., 2004). Several species that maintain high populations within a limited geographical area are reported to harbour C. parvum (Bednarska et al., 2003; Atwill et al., 2004; Ziegler et al., 2007). Environmental loading of Cryptosporidium oocysts depends upon a variety of factors, including the number of infected hosts within a geographical region (Slifko et al., 2000). As hosts these animals constitute an uncontrolled source of pathogen pollution contributing to the degradation of watershed quality. Contamination of the environment by these populations is possible through excretion and through the process of manure-spreading on farmland (Barwick et al., 2003). Runoff from the land contaminated by these sources may serve as a vehicle through which the Cryptosporidium oocysts can be transported into water sources. In this study we carried out a quantitative risk assessment analysis to evaluate the potential risk that dairy herds and wildlife, which resides in the watershed ecosystem, pose to the water supply system, with emphasis at the reservoir level.

Materials and Methods A quantitative risk assessment approach using a pathway model was used to address the stated objective. The risk assessment approach is a systematic process

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that involves the collection, collation and integration of information on a specific hazard to assess its adverse consequences on a particular population (Covello and Merkhofer, 1993). The emphasis in this study was on the release and exposure assessments of C. parvum from components of the watershed ecosystem. The study focused on the New York City and Upper Susquehanna watersheds. Information relating to the likelihood of shedding zoonotic C. parvum and the number of oocysts shed by dairy animals was collected using a couple of crosssectional studies and a longitudinal study (Wade et al., 2000; Starkey et al., 2005, 2006). In addition, data on risk factors hypothesized to associate with the presence of these genotypes in dairy populations in these two watersheds were also collected and analysed (Mohammed et al., 1999; Nydam et al., 2001; Starkey et al., 2005, 2006). In addition to studies on dairy animals, we carried out a cross-sectional study to determine the spectrum of wildlife species that shed C. parvum in the watershed ecosystem (Ziegler et al., 2007). Data on the amount of oocysts shed by each species and the factors that were associated with the presence of the protozoan in these animals were also collected. To complement our understanding of the dynamics of this hazard in the watershed ecosystem we carried out a study to determine the presence of C. parvum in the soil and associated factors (Barwick et al., 2003). We developed a scenario delineating the pathway by which the hazard would reach the stream edge in the hope of capturing the adverse effects associated with the presence of C. parvum in the watershed ecosystem. Figure 10.1 shows the conceptual framework for the fault-tree pathway model used in this study. The primary sources of oocysts were dairy farms, wildlife and agricultural land. These are represented by circles in the figure. Agricultural land was included in the pathway as a separate component because the land could be fertilized by sludge from other sources outside the watershed system, e.g. sludge from neighbouring cities. Since the oocysts are not motile they can either be transported mechanically from the immediate environment of the calf to different sites or moved with the manure as it is carried in waterbodies or runoffs. Only new manure was assumed to be stored near the barn area. Manure that was stored or spread on the agricultural fields had the potential to contaminate the reservoir with oocysts through runoff from the fields. A proportion of the manure produced daily by calves housed in barns, animal houses or the farmyard would also be washed off towards the reservoir. Our studies and others have shown that calves under 30 days of age are at high risk of infection with C. parvum (Wade et al., 2000; Starkey et al., 2005). Figure 10.2 shows the pattern of infection among calves enrolled in the longitudinal study as the data were analysed using the survival analysis method (Starkey et al., 2005). The median time to infection among the calves younger than 60 days of age was 15 days. The number of oocysts shed daily on each farm depends on the number of animals at risk, the infection rate within the farm, and the amount of faeces produced by each calf (Nydam et al., 2005). To capture the potential uncertainty in the estimates of these parameters at the watershed level, the respective distributions were assumed using the parameters obtained through our studies (Table 10.1). The quantity of faeces produced by each calf varies

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Top event

Risk of C. parvum

AND gate

Others

Agricultural fields

OR gate

Wildlife

Species

Sources

Other sources

Cattle

Calves

Soil

Fig. 10.1. The fault-tree scenario describing the pathway by which C. parvum could be transported from the sources to the reservoir.

Time of infection

Survivorship S(t)

1.00

0.75

0.50

0.25

0.00 0

20

40

60

Follow-up time (days)

Fig. 10.2. Time to infection among calves in the longitudinal study analysed using the Kaplan-Meier method.

daily and we assumed that this variability could be captured using a Poisson distribution (Wilkerson et al., 1997). Other parameters that were used in the model are described in Table 10.1. Isolates recovered from dairy cattle in these two watersheds were genotyped using the methods described in our previous study (Starkey et al., 2006). Only the C. parvum genotype, which represented

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Table 10.1. Description, distribution of input variables and the source of the estimates used in the analysis. Variable description Dairy animals Number of herds Number of stock per herd Proportion of calves in a herd Shedding rate per calf Amount of faeces per day Oocysts per gram per day Efficiency of composting pH of the soil Efficiency of soil neutralization Wildlife Occurrence in wildlife Amount of faeces per animal Oocysts per gram per day Cryptosporidium in agricultural field Occurrence in soil Oocysts per gram

Value/Parameter

Source/Reference

Pert (109, 258, 400) Normal (100, 20) Pert (0.01,0.03,0.05) Pert (0.02, 0.08, 0.12) Poisson (1) Exponent (5.6) Pert (0.2, 0.5, 0.7) Triangular (6, 7.4, 8.6) Binomial (0.5, 200)

Barwick et al. (2003) Starkey et al. (2006)

Beta (409, 2992) Lognormal (4, 1) Exponent (3)

Ziegler et al. (2007) Atwill et al. (2004)

Beta (133, 647) Pert (10, 100, 1000)

Barwick et al. (2003) Barwick et al. (2003)

Starkey et al. (2005) Wilkerson et al. (1997) Nydam et al. (2005) Kato et al. (2003) Barwick et al. (2003) Barwick et al. (2003)

61% of all of the isolates that were recovered from cattle in the watershed, was used in the analysis. Some of the practices for managing manure include composting, storage until spreading, or daily spreading on agricultural fields. We assumed that composting is 100% efficacious. During storage and spreading there was some loss due to desiccation of the oocysts as a result of high temperature or dryness (Robertson et al., 1992; Kato et al., 2001; Barwick et al., 2003). In addition to dairy calves we considered the potential contribution from wildlife and agricultural fields. An estimate of the prevalence among the wildlife population in the watershed ecosystem was obtained from our study (Ziegler et al., 2007). An average estimate of 12% for all wildlife in the study was used in the analysis (Table 10.1). To capture the potential variability in this estimate, we used the beta distribution. We observed that the average number of oocysts shed per animal was 1000/g of faeces. An estimate of the average amount of faeces produced by the wildlife was obtained from Atwill et al. (2004). Oocysts were recovered from agricultural land at a rate of 17%, and each gram contained on average 100 oocysts (Barwick et al., 2003). These estimates were used in the final analysis with the respective assumed distribution (Table 10.1). To capture the variability and uncertainty associated with some of the parameters used in the simulation, we assumed that these estimates were obtained from theoretical distributions. We ran the simulation with respective parameters using @risk software (Palisade Corporation, Ithaca, New York, USA).

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Results and Discussion The results of the simulation are shown in Fig. 10.3. Our analysis, using the assumptions about the number of affected calves, the size of the herd, the decay and the wash-off rates, and the presence of oocysts in wildlife and in agricultural fields, showed that on average 10 oocysts could be detected per litre of water per day in the reservoir. However, in the majority of cases it was unlikely that oocysts would be detected in the water (probability 0.078). In a few instances the analysis showed that it would be likely to find more than 24 oocysts per litre in the water samples collected from the reservoir. Although this value was within the range reported in the literature, the associated probability of finding such a concentration was less than 1% (Gibson et al., 1998). This observed number is consistent with what has been reported in the literature. The results reported in this study were similar to the findings of Okun et al. (1997). These authors reported on the average concentration of Cryptosporidium oocysts at the Kensico Reservoir which feeds into the New York City supply system. In our sensitivity analysis several factors influenced the likelihood of finding oocysts at the reservoir. These factors included the level of infection on the farm in general, the shedding rate by the calves, and the pH of the soil. Modification of these practices is likely to reduce the output or environmental loading from dairy farms. In our analysis we assumed that the recovery rate of the water sampling methods was 100%. Methods used in water sampling to date have a recovery rate much lower than that. Our analysis, despite its limitations, has demonstrated the potential contributory role of dairy farms to the risk of water degradation by zoonotic genotypes of Cryptosporidium, namely C. parvum. However, this risk is likely to be mitigated by the implementation of routine practices at the farm level, such as the use of milk replacer, improving the biosecurity measures, or the use of salt to neutralize the soil. 0.100 0.090

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Fig. 10.3. Results of the simulation of the average number of C. parvum oocysts per litre of water per day.

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References Appelbee, A.J., Thompson, R. and Olson, M.E. (2005) Giardia and Cryptosporidium in mammalian wildlife: current status and future needs. Trends in Parasitology 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). Applied and Environmental Microbiology 70, 6748–6752. Barwick, R.S., Mohammed, H.O., White, M.E. and Bryant, R.B. (2003) Factors associated with the likelihood of Giardia spp. and Cryptosporidium spp. in soil from dairy farms. Journal of Dairy Science 86, 784–791. Bednarska, M., Bajer, A., Kulis, K. and Sinski, E. (2003) Biological characterisation of Cryptosporidium parvum isolates of wildlife rodents in Poland. Annals of Agricultural and Environmental Medicine 10, 163–169. Daszak, P., Cunningham, A.A. and Hyatt, A.D. (2000) Emerging infectious diseases of wildlife: threats to biodiversity and human health. Science 287, 443–449. Fayer, R., Speer, C.A. and Dubey, J.P. (1997) The general biology of Cryptosporidium. In: Fayer, R. (ed.) Cryptosporidium and Cryptosporidiosis. CRC Press, Boca Raton, FL, pp. 1–42. Fayer, R., Santín, M. and Xiao, L. (2005) Cryptosporidium bovis n. sp. (Apicomplexa: Cryptosporidiidae) in cattle (Bos taurus). Journal of Parasitology 91, 624–629. Garber, L.P., Salman, M.D., Hurd, H.S., Keefe, T. and Schlater, J.L. (1994) Potential risk factors for Cryptosporidium infection in dairy calves. Journal of the American Veterinary Medical Association 205, 86–91. Hunter, P.R. and Thompson, R.C.A. (2005) The zoonotic transmission of Giardia and Cryptosporidium. International Journal for Parasitology 35, 1181–1190. 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. Veterinary Parasitology 11, 153–159. Kruse, H., Kirkemo, A.M. and Handeland, K. (2004) Wildlife as source of zoonotic infections. Emerging Infectious Diseases 10, 2067–2072. Nydam, D.V., Lindergard, G., Santucci, F., Schaaf, S.L., Wade, S.E. and Mohammed, H.O. (2005) Risk of infection with Cryptosporidium parvum and Cryptosporidium hominis in dairy cattle in the New York City watershed. American Journal of Veterinary Research 66, 413–417. O’Handley, R.M. and Olson, M.E. (2006) Giardiasis and cryptosporidiosis in ruminants. Veterinary Clinics of North America: Food Animal Practice 22, 623–643. Olson, M.E., Thorlakson, C.L., Deselliers, L., Morck, D.W. and McAllister, T.A. (1997) Giardia and Cryptosporidium in Canadian farm animals. Veterinary Parasitology 68, 375–381. Robertson, L.J., Campbell, A.T. and Smith, H.V. (1992) Survival of Cryptosporidium parvum oocysts under various environmental pressures. Applied and Environmental Microbiology 58, 3494–3500. 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. Veterinary Parasitology 122, 103–117. Shukla, R., Giraldo, P., Kraliz, A., Finnigan, M. and Sanchez, A.L. (2006) Cryptosporidium spp. and other zoonotic enteric parasites in a sample of domestic dogs and cats in the Niagara region of Ontario. Canadian Veterinary Journal 47, 1179–1184.

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11

Clinical Presentation in Cryptosporidium-infected Patients

L.M. KORTBEEK National Institute for Public Health and the Environment (RIVM), Bilthoven, The Netherlands

Abstract In immunocompetent patients Cryptosporidium can lead to a self-limited diarrhoea, sometimes recurrent. Other symptoms are vomiting, nausea, decreased appetite, weight loss, flatulence and abdominal pain and cramps. High levels of oocyst shedding have been associated with inflammation, villous atrophy and subsequent malabsorption. In immunocompromised patients CD4 T cell counts at <200 mm3 are associated with persistent (>30 days) diarrhoeal infection and can lead to severe illness. The knowledge of the clinical presentation of Cryptosporidium is changing since it is now possible to type the different strains. In C. hominis cases, non-gastrointestinal symptoms (e.g. joint pain, eye pain, headache, dizziness and fatigue) were seen more often than in cases of C. parvum. In young children, infections with C. hominis and, if symptomatic, C. parvum, are often heavy, associated with faecal lactoferrin and growth shortfalls. C. hominis appears to stimulate inflammation irrespective of age; this raises important questions regarding how it may be specifically inducing greater pro-inflammatory responses. The role of immunity was studied in healthy volunteers. The results indicate that prior exposure to C. parvum provides protection from infection and illness at low oocyst doses. Cryptosporidium can be a cause of serious disease, with a spectrum of symptoms that can vary between asymptomatic infection (non-diarrhoeal but not healthy) to serious infection causing death. The importance of extra-intestinal symptoms is not yet clear and needs to be studied. Typing and subtyping of Cryptosporidium in a standardized way will probably change our ideas and knowledge of clinical symptomatology of cryptosporidiosis in the near future.

Introduction Although Cryptosporidium has been known as an animal parasite since first being described by Tyzzer in 1907, it was not recognized as a human parasite until 1976. The first report of Cryptosporidium as the causative agent of diarrhoea in a 3-year-old child with self-limiting enterocolitis was by Nime et al. (1976). In the same year, Meisel et al. published a paper which reported that Cryptosporidium © CAB International 2009. Giardia and Cryptosporidium: From Molecules to Disease (eds G. Ortega-Pierres et al.)

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had been found in an immunosuppressed patient (Meisel et al., 1976). It was, however, not until the start of the HIV epidemic in the 1980s that Cryptosporidium was recognized as an important human parasite (Petersen, 1992).

Clinical Symptoms The clinical symptoms of Cryptosporidium differ between non-immunocompromised and immunocompromised individuals.

Non-immunocompromised individuals In non-immunocompromised individuals, the disease is usually a self-limiting acute gastroenteritis, characterized by vomiting, weight loss, fever, watery diarrhoea, cramping, abdominal pains, flatulence, malaise and myalgia (Chin, 2000). Recurrent gastrointestinal symptoms may occur in 30–40% of the cases. The duration of the symptoms is about 13 days but this can vary between 1 day and more than 26 days. The severity of the infection is related to the condition and age of the patient. In young children, an important factor is dehydration, especially in children living in developing countries. In a study of 191 children with C. parvum in Uganda, 12.6% died, compared with 6.2% for children without C. parvum in Uganda (Tumwine et al., 2003). Immunocompetent patients do not usually require medical treatment. The number of drugs that are effective is very limited. Most anti-diarrhoeal drugs, such as paramomycin, have no proven effect. In a study in Egypt, Rossignol et al. (2006) demonstrated an effect of a 3-day course of 2 dd 500 mg nitazoxanide, which significantly improved the resolution of diarrhoea and parasitological eradication in non-immunodeficient patients aged 12 years and over. Nitazoxanide has been FDA approved and was marketed in the USA in 2003 (Hussar, 2004).

Immunocompromised individuals In immunosuppressed individuals, i.e. HIV-infected patients with low CD4 T cell counts (<200 mm3), patients undergoing chemotherapy, elderly people and transplant patients, among others, the disease can be severe and debilitating. Patients with idiopathic diarrhoea, biliary disease, primary T cell immune deficiency, acute leukaemia, lymphoma or bone marrow transplant cases, and to a lesser extent those with solid organ transplants and patients taking T cell immunosuppressants, are at high risk for cryptosporidiosis. These patients may develop severe chronic diarrhoeal disease or atypical gastrointestinal disease such as cholangitis, cholecystitis and hepatitis (R.M. Chalmers, 2007, Morelia, personal communication). Infection is associated with persistent diarrhoeal infection, lasting more than 30 days, and can also result in pulmonary or tracheal cryptosporidiosis, low-grade fever or severe intestinal symptoms.

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The most effective therapy in patients with AIDS is highly active antiretroviral treatment (HAART) of HIV. A Cochrane study to assess the efficacy of interventions for the treatment and prevention of cryptosporidiosis among immunocompromised individuals confirms the absence of evidence for effective agents in the management of cryptosporidiosis. The results of this study indicate that nitaxozanide reduces the load of parasites and may be useful in immunocompetent individuals. Due to the seriousness of the potential outcomes of cryptosporidiosis, the use of nitaxozanide should also be considered in immunocompromised patients (Abubakar et al., 2007). The natural history of cryptosporidiosis and its relationship with immune response has been described by Pozio et al. (1997) in their study of a waterborne outbreak in Italy in 1995–1997 involving members of a drug rehabilitation community, their children, and staff members. Of this community, 19.6% were HIVpositive. The attack rate of clinical cryptosporidiosis was 13.6% for HIV-negative members of the community and 30.7% for HIV-positive individuals. The CD4 cell count was indirectly proportional to the attack rate. In general terms, the severity of symptoms was greater among HIV-positive individuals and among those with lower CD4 cell counts. The effect was particularly clear for those with <100 CD4 cells/mm3. Other clinical symptoms, such as abdominal cramps, nausea, vomiting, flatulence and fever, were seen in both groups, but most of them more often in the HIV-positive group. None of the 190 HIV-negative people who complained of diarrhoea at the end of January or at the beginning of February had recurrent or chronic disease. Diarrhoea lasting longer than 14 days was observed only in 22 HIV-positive individuals who had ≤ 300 CD4 cells/mm3. For six of these, the diarrhoea lasted ≥ 30 days. Sera samples of 198 individuals, taken before and after the outbreak, were available. Twenty-eight (14.1%) of the 198 serum samples were already positive (titre 1/50) for specific IgG versus C. parvum antigen before the outbreak. Eightyseven of the 170 initially seronegative individuals (51.2%) developed specific antibodies during the outbreak, reaching titres up to 1/200. As suggested by the attack rates, only a low percentage (17.1%) of the individuals exposed to Cryptosporidium infection developed clinical symptoms, while antibodies against the parasite were detected in 58.0% of the systematic sample. According to these findings, since 17.1% of the participants developed diarrhoea, the authors roughly estimated that almost 40% of infected individuals developed clinical disease. This observation is supported by the rate of seroconversion (52.0%) in asymptomatic HIV-negative individuals, which certainly underestimates the ill-to-infected ratio, since not all infected individuals had detectable specific antibodies in their serum samples collected in March and April, shown by the percentage of seroconversion (90.5%) in symptomatic HIV-negative individuals (Pozio et al., 1997).

Molecular Typing Our knowledge of the clinical presentation of Cryptosporidium is changing, since it is now possible to type the different strains. In the UK, Hunter et al. (2004)

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showed a clear difference in the symptomatology of sporadic cases of cryptosporidiosis between C. hominis and C. parvum. They observed the frequent recurrence of gastrointestinal-related symptoms (e.g. loss of appetite, recurrent vomiting, abdominal pain and diarrhoea) reported by people after recovery from infections caused both by C. parvum and by C. hominis. There were, however, significant differences in the occurrence of non-gastrointestinal symptoms (e.g. joint pain, eye pain, headache, dizziness and fatigue) between the two groups. Following infections with C. hominis, 44.3% had at least one non-gastrointestinal symptom and 27.9% had two or more of these symptoms. This was lower in C. parvum infections, respectively 28.0% and 4.0%, and in a control group (15.1% and 5.2%, respectively) (Hunter et al., 2004). In a study of children in Peru, V. Cama showed an association of different symptoms with infection caused by different subtypes. Cryptosporidium-positive samples were subtyped using the GP60 gene. Subtype C. hominis Ib was associated with nausea, vomiting and general malaise, while subtypes Ia, Ib and Ie were associated with diarrhoea (V. Cama, 2007, Morelia, personal communication).

Asymptomatic infections Asymptomatic infections can now also be studied in more detail. In a study of dairy farm workers and their household contacts in Zimbabwe, 18 asymptomatic Cryptosporidium cases were detected. Cryptosporidium parvum was detected in 12 samples (66.7%; 75% of the positive farm workers and in 60% of the household contacts) and C. hominis in 3 samples (16.7%), while the dairy harboured C. parvum (66%) and C. bovis 33% (Siwila et al., 2007). Asymptomatic infections can also be seen in HIV-positive patients. This seems to occur more frequently with C. hominis than with C. parvum. In a study of HIV-positive adults in Tanzania, Houpt et al. (2005) showed that C. hominis was associated with a longer duration of symptoms, a higher rate of asymptomatic infection, and a lower CD4 cell count compared with C. parvum-infected patients (P < 0.05). This study suggests there may be important differences in the natural history of Cryptosporidium infection in HIV-infected people, depending on parasite species (Houpt et al., 2005). Using more sensitive methods to detect Cryptosporidium, such as PCR, more asymptomatic cases will probably be detected in future. However, it remains to be seen whether these cases are really asymptomatic, or only lacking diarrhoea. There is only limited information on growth and mental development in asymptomatic cases. In a study in Brazil, Bushen et al. (2007) showed that, even in asymptomatic children, C. hominis is associated with heavy infection, lactoferrin and growth shortfalls which are more significant than previously thought. Faecal lactoferrin is a sensitive marker for faecal leucocytes and hence an inflammatory process. It was found that 68% of the Cryptosporidium-infected children were lactoferrin-positive, with no relationship to Cryptosporidium genotype. High levels of oocyst shedding were associated with inflammation, villous atrophy and subsequent malabsorption. In symptomatic C. parvum cases, the numbers of oocysts shed were higher than in asymptomatic cases. Asymptomatic C.

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hominis cases had high numbers of oocysts and long-term growth shortfall; half were lactoferrin-positive (Bushen et al., 2007).

Immunity Another issue that needs attention is the role of immunity in the clinical presentation of Cryptosporidium infections. Chappell et al. (1999) showed, in a study in healthy volunteers, that there is a significant difference in the number of oocysts that are required to cause an infection: the ID50 was 1880 oocysts in healthy volunteers with pre-existing a-C. parvum serum IgG antibodies, while the ID50 was 83 oocysts in volunteers without pre-existing a-C. parvum serum IgG. These data indicate that prior exposure to C. parvum provides protection from infection and illness at low oocyst doses (Chappell et al., 1999). In another study in healthy volunteers it was shown that there is a difference in the serological response by the different Cryptosporidium types. In contrast to C. parvum (Iowa isolate), C. hominis resulted in a serological response in 8 (38.1%) of 21 challenged volunteers and in 8 (53.3%) of 15 who had evidence of infection. Interestingly, only volunteers receiving 30 oocysts or more had a serum IgG response, even though all had diarrhoea or faecal oocyst shedding. Furthermore, the degree of response was influenced by post-challenge outcome. Volunteers who had a diarrhoeal illness and who shed detectable levels of oocysts yielded the highest responses (Chappell et al., 2006).

Where Does the Science Go from Here? One of the questions that was asked at the II International Giardia and Cryptosporidium Conference held in May 2007 in Morelia, Mexico, was: ‘Where should the science go from here?’ A paper published after the first Giardia and Cryptosporidium meeting in Amsterdam (2003) by Savioli et al. put Giardia and Cryptosporidium on the list of neglected diseases. Most of the important questions that were raised then still need to be addressed (Savioli et al., 2006). In order to understand the clinical significance of cryptosporidiosis, it is important to study the sequelae of an infection and the mechanisms behind these sequelae. We should also study the frequency of the non-intestinal symptoms of the infection (arthritis, uveitis and urticaria) and determine their severity: the same applies to the following unanswered questions: ● ● ● ● ● ●

How often does malabsorption occur in cryptosporidiosis? What are the main risk groups? Which markers can be used to detect it? How often do deficiencies of micronutrients (vitamin A, minerals, etc.) occur? Is there a relation with post-infection IBS/IBD? If yes, how frequently does this occur and what are the risk groups?

To be able to compare studies in different parts of the world and in different settings, we need definitions for clinical cryptosporidiosis and giardiasis; and also

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a definition for diarrhoea (e.g. ≥ 3 stools/day and different consistency). In addition, we need to establish the clinical significance of laboratory-confirmed cryptosporidiosis and giardiasis: the significance of an isolated positive PCR without detectable cysts or oocysts; whether we need multiple sampling; and the value of serum antibodies, etc. We also need to be able to agree on the definition of acute and chronic diarrhoea. I suggest that acute diarrhoea lasts between 1 and 14 days and chronic diarrhoea persists for ≥ 30 days. We need good diagnostic routines so that clinicians know when to test for Giardia or Cryptosporidium. In a normal diagnostic setting it is possible to use available information (age, season, duration of symptoms) to reduce the requests to test for different microorganisms considerably. The methods chosen, or the combination of methods, can also assist in more sensitive and cost-effective diagnostics.

Conclusions Cryptosporidium can cause serious disease with a spectrum of symptoms that can vary between asymptomatic infection (non-diarrhoeal but not healthy) to serious infection causing death. The importance of extra-intestinal symptoms is not yet clear and needs to be studied. Typing and subtyping of Cryptosporidium in a standardized way will probably change our ideas and knowledge of clinical symptomatology of cryptosporidiosis in the near future.

References Abubakar, I., Aliyu, S.H., Arumugam, C., Hunter, P.R. and Usman, N.K. (2007) Prevention and treatment of cryptosporidiosis in immunocompromised patients. Cochrane Database Systematic Reviews 24, CD004932. 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. Transactions of the Royal Society of Tropical Medicine and Hygiene 101, 378–384. Chappell, C.L., Okhuysen, P.C., Sterling, C.R., Wang, C., Jakubowski, W., Dupont, H.L. (1999) Infectivity of Cryptosporidium parvum in healthy adults with pre-existing anti-C. parvum serum immunoglobulin. American Journal of Tropical Medicine and Hygiene 60, 157–164. 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. American Journal of Tropical Medicine and Hygiene 75, 851–857. Chin, J. (2000) Cryptosporidiosis. In: Chin, J. (ed) Control of Communicable Diseases Manual, 17th edn. American Public Health Association, Washington, DC, pp. 134–137. 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. American Journal of Tropical Medicine and Hygiene 73, 520–522.

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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. Clinical Infectious Diseases 39, 504–510. Hussar, D.A. (2004) New drugs of 2003. Journal of the American Pharmaceutical Association 44, 168–206. 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. 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. Petersen, C. (1992) Cryptosporidiosis in patients infected with the human immunodeficiency virus. Clinical Infectious Diseases 15, 903–909. Pozio, E., Rezza, G., Boschini, A., Pezzotti, P., Tamburrini, A., Rossi, P., Di Fine, M., Smacchia, C., Schiesari, A., Gattei, E., Zucconi, R. and Ballarini, P. (1997) Clinical cryptosporidiosis and human immunodeficiency virus (HIV)–Induced immunosuppression: findings from a longitudinal study of HIV-positive and HIV-negative former injection drug users. Journal of Infectious Diseases 176, 969–975. Rossignol, J.-F., Kabil, S.M., El-Gohary, Y. and Younis, A.M. (2006) Effect of nitazoxanide in diarrhea and enteritis caused by Cryptosporidium species. Clinical Gastroenterology and Hepatology 4, 320–324. Savioli, L., Smith, H. and Thompson, A. (2006) Giardia and Cryptosporidium join the ‘Neglected Diseases Initiative’. Trends in Parasitology 22, 203–208. Siwila, J., Phiri, I.G., Vercruysse, J., Goma, F., Gabriel, S., Claerebout, E. and Geurden, T. (2007) Asymptomatic cryptosporidiosis in Zambian dairy farm workers and their household members. Transactions of the Royal Society of Tropical Medicine and Hygiene 101, 733–734. 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. American Journal of Tropical Medicine and Hygiene 68, 710–715.

12

Molecular Epidemiology of Cryptosporidium and Giardia Infections

P.R. HUNTER University of East Anglia, Norwich, UK

Abstract For infectious diseases, molecular epidemiology can be defined as the use of molecular (predominantly genetic) methods to distinguish between strains of a microbial pathogen in order to identify markers of virulence or host range or to elucidate different transmission pathways. In this regard, molecular epidemiological methods have the same objectives as other typing methods that have been in use for several decades. The characteristics of a good typing method are typability, reproducibility and discriminatory power. There have been significant advances in typing methods for Cryptosporidium spp. in recent years. The first advances were for methods to distinguish the different species, predominantly C. parvum from C. hominis. More recently, microsatellite markers have been used to distinguish different strains within these two species. Microsatellite typing of C. parvum is sufficiently discriminatory to be useful, although more information is needed to determine reproducibility. Typing methods for C. hominis are not yet sufficiently discriminatory for general use. Molecular typing methods for Giardia are not yet well developed and have not been used in well-designed epidemiological studies.

Introduction The phrase ‘molecular epidemiology’ has become very popular in the title of papers and grant applications in the last few years (Fig. 12.1). So it is perhaps appropriate to discuss what we mean by this before moving on to discuss the molecular epidemiology of Cryptosporidium and Giardia infections. Molecular epidemiology is more generally used to refer to any study that uses molecular methods (and increasingly this means genetic methods) to identify markers of susceptibility in hosts or virulence in pathogens. The other main use of molecular epidemiology is to track the spread of pathogens in order to elucidate transmission pathways. In this regard molecular epidemiology does not differ in its objectives, or indeed its philosophy, from other work on microbial epidemiology that uses non-molecular typing methods. 138

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Any discussion of molecular epidemiology needs to focus on two key areas. The first concerns the methods used to characterize the strains, and the second concerns the epidemiological studies themselves. There seems to be a view in some quarters that because molecular methods have been used then the results are somehow more compelling than those obtained by traditional typing methods. However, the need for proper validation of molecular methods is just as great as that for non-molecular methods. It is somewhat disappointing that proper validation is often not done. This chapter will first discuss the validation of typing methods. Before taking the discussion any further we need to ask what microbiological typing methods are used for. The primary role of a typing method is to distinguish between two strains. No typing method can say that two strains are the same, only that they are different. For example, if serotyping of two strains of E. coli shows that both are E. coli O157 this does not necessarily imply that they are epidemiologically related. On the other hand, if one is O157 and the other is O135 we can be confident that the two are not the same. The other use of typing methods is in the identification of markers that may indicate particular virulence, host preferences, or routes of transmission. In the E. coli serotyping example, if a strain of E. coli O157 was isolated from a patient with bloody diarrhoea, then we would be fairly confident that this strain was an enterohaemorrhagic strain. However, one should not make the mistake of assuming that the marker = the virulence factor. Many strains of E. coli O157 are not particularly pathogenic and, if one was isolated from an environmental water sample, it would be dangerous to assume that it was enterohaemorrhagic without tests to identify the virulence factor directly. It is also important to understand the differences between typing methods and typing schemes (Hunter, 1991). A typing method is any method that is

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used to distinguish between different strains of the same species. A typing scheme, on the other hand, is a typing method that has been subjected to rigorous inter-laboratory comparisons and has been agreed on as an international standard. Examples include the phage typing of Staphylococcus aureus and serotyping of Salmonella or E. coli. There are six characteristics of a typing method that need to be considered when assessing its value (Hunter, 1991): 1. 2. 3. 4. 5. 6.

Typability. Reproducibility. Discriminatory power. Cost. Timeliness. Ease of use.

The latter three characteristics are fairly obvious. An ideal typing method would be cheap, give a quick result and be sufficiently easy to do that extensive training of staff would not be required. The other characteristics need more explanation. Typability is a measure of the proportion of strains that can be typed by the method. Some strains, for example, simply may not amplify with one of the primers and so cannot be allocated to a specific type (Hunter et al., 2007). Typability is represented as the percentage of strains that can be fully typed by the method. One should be careful not to assume that the non-typing strain is in any way different from a strain that is fully typed. A non-typed strain cannot be said to be different from one that is typed. There are two aspects of reproducibility: in vitro and in vivo reproducibility. In vitro reproducibility is the proportion of strains that, should they be retyped, would give exactly the same result. In vitro reproducibility is primarily a measure of the reproducibility of the typing method itself. Sources of poor in vitro reproducibility include poor operator experience, difficulties with the method itself, and sloppiness in recording results. In vivo reproducibility is a more difficult concept, but is concerned with the stability of the organism itself. Some pathogens have a habit of switching their phenotypes (see Jones et al., 1994). Of particular concern to genetic-based typing methods is the possibility of mutation or, for the case of microsatellites, slippage (e.g. Eckert et al., 2002; Tanriverdi and Widmer, 2006). Whatever the mechanism, if genetic change occurs, then the possibility of getting different type results increases when strains are, in fact, related. Discriminatory power is a measure of the ability of the typing method to distinguish between unrelated strains. Discriminatory power is a function of the number of different stain types identifiable and also the distribution of strains between these different types. Discriminatory power is greater where there are more types, and strains are evenly distributed between types. Poor discriminatory typing methods identify few types, and most strains belong to a single type. In 1988 we proposed an index of discriminatory power that has become the standard way of describing the discriminatory power (Hunter and Gaston 1988). A short while later we described a generalization of the index that is applicable to situations where reproducibility is less than 100% (Hunter, 1990). Other workers

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have described an approach to determine the confidence intervals of estimated discriminatory power. Clearly, discriminatory power will increase as more typing variables are used. One of the uses to which indices of discriminatory power can be put is the choice of parsimonious collections of typing characters or methods to give the most efficient discriminatory tests (Gaston and Hunter, 1989). If one starts with the most discriminatory test, then as each furthermost discriminatory parameter is included, the combined discriminatory power increases, but very soon any increase in discrimination will level off (Fig. 12.2). On the other hand, as the number of variables increases, reproducibility continues to decline. Consequently, increasing the number of variables beyond a certain point is counterproductive.

Initial Work on Cryptosporidium One of the early studies of genotyping and cryptosporidiosis to suggest two distinct genotype types with different host ranges was that of Peng et al. (1997). Based on typing 39 strains from human and animal sources, they established that C. parvum genotype 1 was isolated only from humans whilst genotype 2 was isolated from humans and animals. Furthermore, human cases with genotype 2 tended to report contact with cattle or cattle products. The author of this chapter usually has serious concerns about the strength of conclusions based on the examination of a relatively small number of strains. However, on this occasion the conclusions appear to be sound. McLaughlin et al. (2000) reported a much larger study from the UK. In this study the authors examined 1705 positive faecal samples from humans and 105 from livestock animals. Genotype 1 was detected in 37.8% of human samples, whilst genotype 2(B) was detected in 61.5%. C. meleagridis was detected in 0.3%. On the other hand, all the samples from livestock yielded genotype 2. 1 0.9 0.8

D or R

0.7 0.6

D

0.5

R

0.4 0.3 0.2 0.1 0 1

2

3

4

5

6

7

8

9

10

Number of typing variables

Fig. 12.2. Indication of the impact of increasing numbers of typing variables on discriminatory power (D) and reproducibility (R).

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Probably the first large-scale analytical epidemiological study done on sporadic cryptosporidiosis using these ‘genotyping’ methods was that of Hunter et al. (2004a). In this study the authors were able to identify personal risk factors between genotype 1 and genotype 2 strains. The authors found that the strongest risk factors for genotype 1 or C. hominis were travel abroad (OR = 6.841, 95% CI 2.622, 17.85) and changing nappies (OR = 3.991, 95% CI 1.848, 8.618). By contrast the strongest association for genotype 2 or C. parvum was direct contact with farm animals (OR = 2.653, 95% CI 1.113, 6.323). Of particular interest was the observation that the consumption of tomatoes (OR = 0.317, 95% CI 0.140, 0.719) or other raw vegetables (OR = 0.222, 95% CI 0.086, 0.572) were negatively associated with risk. Using the same population, the authors were also able to show that long-term sequelae were likely to follow infection with C. hominis but not with C. parvum (Hunter et al., 2004b). At one time, these studies would have counted as molecular epidemiological studies. However, since the different genotypes were deemed to be two separate species (Morgan-Ryan et al., 2002), the molecular methods used would be better classified as species identification. Consequently these studies would not now be classed as molecular epidemiology.

Subspecies Typing of Cryptosporidium Most of the currently published studies of strain typing methods for Cryptosporidium spp. have been based on microsatellite markers (Aeillo et al., 1999; Cacciò et al., 2000, 2001; Enemark et al., 2002; Mallon et al., 2003a, 2003b). These studies were generally focused on comparing polymorphisms in human and animal sources. Of particular interest is the work of Mallon et al. (2003a, 2003b), undertaken in Scotland, that identified a cluster of C. parvum strains that were identified in humans but not animals. Based on this finding, they suggested that there was a specific clone of C. parvum that was anthropogenic. This was a valuable insight. However, care must be taken when interpreting such data, especially from relatively small geographical locations and when not accompanied by specific exposure data. In this context the study design would not be able to distinguish between a strain that was truly anthropogenic and one that was zoonotic but the focus of animal infection was localized elsewhere. So, for example, based on just the evidence from this study, it is plausible that the type was a zoonotic infection in the Mediterranean, was acquired by people on holiday and brought back to the UK, and had not yet developed any animal cycle of infection. We have recently published a study that looked at strain typing of both C. parvum and C. hominis using three microsatellite markers: ML1, ML2 and GP60 (Hunter et al., 2007). In this study we typed strains that were part of our earlier case control study (Hunter et al., 2004a). Taken together, these three markers were highly discriminatory for C. parvum (D = 0.957) but very poorly discriminatory for C. hominis (D = 0.197). This means that for C. hominis the method would be able to distinguish between two unrelated strains on fewer than 20% of occasions, and so was not suitable for epidemiological studies of this species.

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The relatively poor typability of C. hominis should come as no surprise. It has been pointed out that, as would be predicted by the theory of adaptive polymorphism, that pathogens with a limited host range and no environmental replication are generally much less diverse than pathogens with a wide host range and/ or environmental life stages (Hunter and Fraser, 1990). For C. parvum, three main clusters were identifiable, one large cluster and two smaller clusters. This was in line with earlier findings of Mallon et al. (2003a, 2003b). All the members of the larger cluster had a single MLT1 fragment length 242 bp while all others were MLT1-227. Interestingly, we were able to show that all cases who reported farm animal contact had the MLT1-242 type, and furthermore the attack rate was more common in rural areas. All C. parvum strains from animals in a related study were also MLT1-242 type. Thus our findings would strengthen Mallon’s suggestion, although again our study would not be able to exclude the hypothesis that MLT1-227 strains were zoonotic in other parts of the world.

Application of Molecular Epidemiology to Outbreak Investigations Microbial typing has a long tradition of use in the investigation of outbreaks of infectious disease. Strain typing data can be helpful in one of several ways. First, typing can be an indicator of the source of infection. For example, as is the case in the investigation of waterborne outbreaks, strain typing may give important indications of the source of contamination. Knowing whether the infection is due to C. parvum or C. hominis will let the investigator know whether or not the source of pollution was human or cattle (e.g. Glaberman et al., 2002; Dalle et al., 2003; Goncalves et al., 2006). Microbial typing is also of value in identifying whether or not individual cases are part of an outbreak. This is important to enhance the strength of the epidemiological studies. If many unrelated cases are included in the analysis, important risk factors may be missed. To date, no good subspecies typing studies of strains from outbreaks of cryptosporidiosis have been reported in the literature. However, preliminary results from the investigation of a waterborne outbreak in the north-west of England have identified multiple MLFT types. The implications of this finding are currently unclear. It may be the case that the outbreaks are in fact due to multiple strains. Alternatively, it may be the case that the typing characters are not sufficiently stable for the investigation of large outbreaks. Whatever the explanation, if it is the case that waterborne outbreaks are usually due to multiple strain types, then the value of strain typing would be severely adversely affected.

Giardia In general, strain typing issues for Giardia are similar to those for Cryptosporidium. However, epidemiological studies for Giardia are not as advanced as those for Cryptosporidium (Hunter and Thompson, 2005). In part, this is because the

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characterization of different species/assemblages is not yet as well accepted as species differentiation in Cryptosporidium. Also it would appear that the methods available for further typing below the species/assemblage level are not as well characterized. One typing method that may be valuable is sequence heterogeneity in the β-giardin gene (Lalle et al., 2005).

Conclusions In recent years the development of molecular typing methods has contributed to our understanding of the epidemiology of cryptosporidiosis and, to a much lesser extent, giardiasis. However, few of the methods have yet to be adequately characterized in the published literature to enable their reproducibility and discriminatory power to be adequately assessed. Furthermore, relatively few of the studies that have used molecular typing methods have actually linked the strain typing data to basic epidemiological data. Many studies have been reported that have simply listed identified strain types in different hosts. Grinberg et al. (2007) have recently pointed out the dangers of relying solely on differences in detected strain types in different host populations to draw too strong a conclusion. There is always a strong desire amongst microbiologists to develop their own tests, and so it is likely that the microbiological community will spend considerable time and effort in developing new typing methods for both Giardia and Cryptosporidium. However, the increase in our knowledge of the epidemiology of these two pathogens will be best served if our community focuses on better characterizing the methods we currently have available. There is also an obvious need for microbiologists and parasitologists who can do strain typing to develop their project proposals in collaboration with epidemiologists. Only then will they really understand what the results of their strain typing are telling us about the epidemiology of these two pathogens.

References Aeillo, A.E., Xiao, L., Limor, J.R., Liu, C., Abrahmason, M.S. and Lal, A.A. (1999) Microsatellite analysis of the human and bovine genotypes of Cryptosporidium parvum. Journal of Eukaryotic Microbiology 46, 46S–47S. Cacciò, 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. Cacciò, S., Spano, F. and Pozio, E. (2001) Large sequence variation at two microsatellite loci among zoonotic (genotype C) isolates of Cryptosporidium parvum. International Journal for Parasitology 31, 1082–1086. 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. Journal of Clinical Microbiology 41, 2690–2693.

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Eckert, K.A., Mowery, A. and Hile, S.E. (2002) Misalignment-mediated DNA polymerase beta mutations: comparison of microsatellite and frame-shift error rates using a forward mutation assay. Biochemistry 41, 10490–10498. 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. Gaston, M.A. and Hunter PR. (1989) Efficient selection of tests for bacteriological typing schemes. Journal of Clinical Pathology 42, 763–766. Glaberman, S., Moore, J.E., Lowery, C.J., Chalmers, R.M., Sulaiman, I., Elwin, K., Rooney, P.J., Millar, B.C., Dooley, J.S.G., Lal, A.A. and Xiao, L. (2002) Three drinkingwater-associated cryptosporidiosis outbreaks, Northern Ireland. Emerging Infectious Diseases 8, 631–633. 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 São Paulo. Clinics (São Paulo, Brazil) 61, 119–126. Grinberg, A., Lopez-Villalobos, N., Pomroy, W., Widmer, G., Smith, H. and Tait, A. (2007) Host-shaped segregation of the Cryptosporidium parvum multilocus genotype repertoire. Epidemiology and Infection 136, 273–278. Hunter, P.R. (1990) Reproducibility and indices of discriminatory power of microbial typing methods. Journal of Clinical Microbiology 28, 1903–1905. Hunter, P.R. (1991) A critical review of typing methods for Candida albicans. Critical Reviews in Microbiology 17, 417–434. Hunter, P.R. and Fraser, C.A.M. (1990) Application of the theory of adaptive polymorphism to the ecology and epidemiology of pathogenic yeasts. Applied and Environmental Microbiology 56, 2219–2222. Hunter, P.R. and Gaston, M.A. (1988) A numerical index of the discriminatory ability of typing systems: an application of Simpson’s index of diversity. Journal of Clinical Microbiology 26, 2465–2566. Hunter, P.R. and Thompson, R.C.A. (2005) The zoonotic transmission of Giardia and Cryptosporidium. International Journal for Parasitology 35, 1181–1190. Hunter, P.R., Hughes, S., Woodhouse, S., Syed, Q., Verlander, N.Q., Chalmers, R.M., Morgan, K., Nichols, G., Beeching, N. and Osborn, K. (2004a) Sporadic cryptosporidiosis case-control study with genotyping. Emerging Infectious Diseases 10, 1241–1249. Hunter, P.R., Hughes, S., Woodhouse, S., Raj, N., Syed, Q., Chalmers, R.M., Verlander, N.Q. and Goodacre, J. (2004b) Health sequelae of human cryptosporidiosis in immunocompetent patients. Clinical Infectious Diseases 39, 504–510. Hunter, P.R., Hadfield, S.J., Wilkinson, D., Lake, I.R., Harrison, F.C.D. and Chalmers, R.M. (2007) Subtypes of Cryptosporidium parvum in humans and disease risk. Emerging Infectious Diseases 13, 82–88. Jones, S., White, G. and Hunter, P.R. (1994) Increased phenotypic switching in strains of Candida albicans associated with invasive infections. Journal of Clinical Microbiology 32, 2869–2870. Lalle, M., Pozio, E., Capelli, G., Bruschi, F., Crotti, D. and Cacciò, S.M. (2005) Genetic heterogeneity at the β-giardin locus among human and animal isolates of Giardia duodenalis and identification of potentially zoonotic subgenotypes. International Journal for Parasitology 35, 207–213. 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. Journal of Molecular Evolution 56, 407–417.

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P.R. Hunter Mallon, M.E., MacLeod, A., Wastling, J.M., Smith, H. and Tait, A. (2003b) Multilocus genotyping of Cryptosporidium parvum Type 2: population genetics and substructuring. Infection Genetics and Evolution 3, 207–218. McLaughlin, 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 1705 fecal samples from humans and 105 fecal samples from livestock animals. Journal of Clinical Microbiology 38, 3984–3990. Morgan-Ryan, U.M., Fall, A., Ward, L.A., Hijjawi, N., Sulaiman, I., Fayer, R., Thompson, R.C.A., Olson, M., Lal, A. and Xiao, L.H. (2002) Cryptosporidium hominis n. sp (Apicomplexa: Cryptosporidiidae) from Homo sapiens. Journal of Eukaryotic Microbiology 49, 433–440. Peng, M.M., Xiao, L., Freeman, A.R., Arrowood, M.J., Escalante, A.A., Weltman, A.C., Ong, C.S.L., Mackenzie, W.R., Lal, A.A. and Beard, C.B. (1997) Genetic polymorphism among Cryptosporidium parvum isolates: evidence of two distinct human transmission cycles. Emerging Infectious Diseases 3, 567–573. Tanriverdi, S. and Widmer, G. (2006) Differential evolution of repetitive sequences in Cryptosporidium parvum and Cryptosporidium hominis. Infection, Genetics and Evolution 6, 113–122.

13

Advances in Diagnosis: is Microscopy Still the Benchmark?

R.M. CHALMERS UK Cryptosporidium Reference Unit, NPHS Microbiology, Swansea, UK

Abstract Despite advances in the biomedical sciences, leading to developments in laboratory diagnostics, examination (usually of faeces) by microscopy remains the keystone of laboratory diagnosis of cryptosporidiosis and giardiasis. In Europe, standard methods include the direct examination of wet preparations (with and without concentration) for Giardia, and tinctorial staining of faecal smears (usually with strong carbol fuchsin) or fluorescent staining (e.g. using Auramine O) for Cryptosporidium. There is an increasing trend in the UK towards the routine use of Auramine O staining, and over 70% of laboratories use this stain, while 26% use a modified Ziehl-Neelsen stain. Immunoassays to detect copro-antigens are gradually being introduced into routine diagnostics, particularly in larger laboratories processing large numbers of samples. These methods are more commonly used in the USA, as is immunofluorescent microscopy, which offers increased sensitivity and specificity over conventional staining. However, cost constraints preclude their use in many countries. Advances in molecular diagnostics have shown that PCR-based methods have exquisite sensitivity in pure (oo)cyst suspensions, but amplification inhibitors and barriers to DNA extraction need to be overcome in faecal samples. Additional benefits are Cryptosporidium species/genotype or Giardia assemblage identification. However, few diagnostic laboratories are well placed to undertake PCR. The modernization and amalgamation of pathology services may be a driver for the increased use of immunoassays and PCR in diagnostic settings, most logically as multiplexed tests for panels of gastrointestinal pathogens. However, while these and other biomedical advances may aid diagnosis, any loss of microscopy skills would be to the detriment of parasitology services worldwide.

Introduction It is over 300 years since Antony van Leeuwenhoek’s microscopic examination of his own stools in 1681 facilitated his description of what we now recognize as Giardia cysts and trophozoites. The first formal description was published by Lambl in 1859, who proposed the name Cercomonas intestinalis. The following © CAB International 2009. Giardia and Cryptosporidium: From Molecules to Disease (eds G. Ortega-Pierres et al.)

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50 years or so saw many proposed genus and species names, until Giardia lamblia and Giardia enterica were proposed for the parasite isolated from humans (Kofoid and Christensen, 1915; Kofoid, 1920). The currently used Giardia duodenalis and Giardia intestinalis are most accurate taxonomically, although some prefer the former since it conforms to Filice’s morphologically based nomenclature published in 1952 (Filice, 1952). Giardia cysts are readily visible in microscopic examination of a wet preparation from a faecal sample, but trophozoites, being more fragile and less likely to survive intact, are seen less frequently. Similarly, astonishingly accurate and detailed early descriptions of Cryptosporidium life cycle stages were made by Ernest E. Tyzzer in 1907, 1910, 1912 and 1929, establishing the life cycle of the parasite. The only element later amended by examination using electron microscopy was the added description of extracellular developmental stages (merozoites and microgametes). Cryptosporidium was initially recognized as an animal pathogen, and staining techniques were improved to detect oocysts in faeces for veterinary diagnosis. Transfer of these improvements to clinical laboratories enabled diagnosis by examination of stools instead of histological staining of gut biopsy. This facilitated important epidemiological studies during the early 1980s, which led to the recognition of the parasite of importance not only as a potentially fatal infection of immunocompromised patients but also as a cause of acute, self-limiting gastroenteritis in the general population, particularly children (Casemore et al., 1985). Widespread testing and reporting of microbiological results to disease surveillance schemes was thus enabled. In England and Wales, this has been ongoing since 1983. The impact of infection with Cryptosporidium and Giardia was recognized by WHO in 2004 by their inclusion in the ‘Neglected Pathogen Initiative’. Routine laboratory diagnosis of these pathogens is undertaken for a variety of reasons: ● ● ● ● ● ●

Routine clinical diagnosis for patient care. For surveillance and public health purposes. Contact screening. Outbreak/cluster investigation. Enhanced surveillance, e.g. incorporating specialist test results or patient data. Special studies, e.g. carriage rates; experimental studies (transmission/infectivity/treatment trials).

This chapter describes the current situation with regard to routine and specialist diagnostic processes for Cryptosporidium and Giardia.

Testing Algorithms and Diagnostic Procedures Testing algorithms describe the appropriate ordering and use of laboratory tests and are designed to provide guidance to ensure appropriate sample submission, reduce unnecessary testing and accelerate the diagnostic process. Expansion of the algorithm may include investigation and management of suspected and confirmed cases of illness and reporting systems. Algorithms may be produced for individual pathogens, symptoms (e.g. gastrointestinal illness), patient groups (e.g. high-risk patients, travellers returning from abroad) or settings (e.g. schools

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and other child care centres, hospitals) at the local, region, national or international level. National diagnostic test algorithms for Cryptosporidium and Giardia are currently produced in the form of guidance, such as the Investigation of Specimens other than Blood for Parasites produced by the Health Protection Agency (HPA) in England (see http://www.hpa-standardmethods.org.uk/documents/ bsop/pdf/bsop31.pdf) and its sister document Staining Procedures (http://www. hpa-standardmethods.org.uk/documents/bsopTP/pdf/bsoptp39.pdf). The equivalent in the USA is Laboratory Detection of Parasites of Public Health Concern produced by the Centers for Disease Control and Prevention (http://www.dpd. cdc.gov/dpdx/HTML/Cryptosporidiosis.htm). UK veterinary testing is guided by a working group for the Harmonization of Laboratory Diagnostic Methods of the National Surveillance Group on Diseases and Infections of Animals for the testing of samples from farmed animals in Veterinary Laboratories Agency (VLA) regional laboratories. The methods used are recognized by the Office International des Epizooties Manual of Diagnostic Tests and Vaccines for Terrestrial Animals, published in 2004 and updated in 2008 (http://www.oie.int/eng/normes/ mmanual/A_summry.htm). However, this document does not include Giardia, the impact of which on livestock and companion animals may be underestimated because in some countries, including the UK, it is not routinely reported by veterinary laboratories. In many countries Cryptosporidium is only tested for on specific request or in HIV & AIDS patients, and Giardia only as part of an ova and parasites (O&P) request. Requesting primary care physicians and clinicians may not be aware that Cryptosporidium is not necessarily included in this. The UK guidance is for all samples from symptomatic patients to be tested for Cryptosporidium, based on the findings of a working group published in 1993 (Casemore and Roberts, 1993). Testing for Giardia often depends on the patient history (mainly recent foreign travel) and clinical details. Recent surveys by the UK Cryptosporidium Reference Unit have shown that there is an increasing trend for compliance in testing for Cryptosporidium (Chalmers et al., 2002; UK Cryptosporidium Reference Unit, unpublished data). In a survey of 187 laboratories carried out in 2003, 53% reported routinely testing all diagnostic stools from community cases for Cryptosporidium, a figure which had risen to 78% by 2006. The overall positivity rate is in the region of 1%. In the 2003 survey, 36% of laboratories tested all such stools by microscopic examination of a wet preparation, which would include examination for Giardia. It is common practice in the UK to treat hospital in-patients differently from community cases and a ‘3- or 5-day rule’ is often applied. This means that if a patient who had been in hospital develops diarrhoeal illness after this time period, they are not tested for the same suite of pathogens as community cases, and tests for Cryptosporidium and Giardia are not performed unless specifically requested. In VLA regional veterinary laboratories, Cryptosporidium is examined for only in neonatal animals with diarrhoea, unless pathological examination (for example post mortem) suggests the involvement of this parasite. Diagnosis of intestinal protozoa traditionally relies on the detection of life cycle stages in appropriate samples. For Cryptosporidium this is mainly the oocyst, and for Giardia the cyst, in faeces (Table 13.1). Giardia trophozoites may

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Table 13.1.

Routine diagnostic procedures for Cryptosporidium in faeces.

Procedure

Detection method

Microscopy

Modified acid-fast stains

Immunoassay

Microscopy

Diagnostic target

Format

Advantages

Disadvantages

Oocyst

Fixed microscopy slide

Low cost

Fluorescent stains

Oocyst

Fixed microscopy slide

Immunofluorescent stains

Oocyst

Fixed microscopy slide

Enzyme

Antigen

Microplate

Chromatographic

Antigen

Strip-based cartridge

Modified acid-fast stains

Oocyst

Fixed microscopy slide

Low cost Improved specificity over acid-fast stains Low power, more rapid screening Improved sensitivity over acid-fast and fluorescent stains Mass testing Lower skill base Lower skill base than microscopy Portable (field testing) See above

Skilled microscopist Detects other pathogens Poor sensitivity and specificity Skilled microscopist

Higher cost

High cost False positives High cost False positives and negatives See above R.M. Chalmers

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also occasionally be seen. Throughout Europe it is current practice for most diagnostic stools to be submitted fresh, without preservatives. While this has some advantages, the vegetative stages of many parasites may degrade if there are delays in processing. Testing of other sample types depends on the clinical history of the patient: for immunocompetent patients the main symptom is diarrhoea and stool is the most common and appropriate specimen type. Other specimen types for diagnosis of Cryptosporidium, such as tissue biopsy, sputum, bronchoalveolar lavage and bile are generally from severely immunocompromised patients who may suffer from extra-intestinal infection or infection of the biliary tree (Farthing, 2000; Hunter and Nichols, 2002). Diagnosis in cases of suspected biliary involvement may also be assisted by imaging techniques, including ultrasound, CT scans and, more specifically, endoscopic retrograde cholangio-pancreatography (the definitive test for disruption of biliary anatomy). Difficulty in detecting Giardia may be assisted by testing duodenal aspirate. Detection methods for Cryptosporidium oocysts are traditionally tinctorial or fluorescent staining of faecal smears examined by bright field or epifluorescence microscopy, respectively (Nichols and Thom, 1984; Casemore, 1991), while Giardia cysts and occasionally trophozoites can be seen in examination of a wet preparation (Table 13.1). Methods recommended in the HPA guidance are stool microscopy, specifically modified Ziehl-Neelsen or auramine phenol staining for Cryptosporidium. These methods are also specified for animal testing in VLA laboratories. Improved sensitivity is provided by immunofluorescence microscopy (IFM) using a fluorescein isothiocyanate (FITC)-labelled monoclonal antibody raised against oocyst wall antigens (Arrowood, 1997; Casemore, 1992), which is also available in combination for the simultaneous detection of these two protozoa. These stains can be applied to unconcentrated as well as concentrated faecal samples. Concentration by the formol-ether method is suitable and there are commercially available kits providing the plastic ware to facilitate this. Garcia (2001) provides an excellent laboratory manual for parasitological diagnosis. In a survey undertaken in 2006 of 170 primary diagnostic laboratories in the UK, 26% used modified Ziehl-Neesen staining and 72% used auramine phenol staining (UK Cryptosporidium Reference Unit, unpublished data). Just three laboratories used other methods: one used flotation and direct examination and two used antigen detection kits. None used IFM as a primary diagnostic test, for reasons of cost, although this is available as a specialist reference test at the UK Cryptosporidium Reference Unit (see below). The emergence of copro-antigen test kit use in the UK is recent: in a previous survey 3 years earlier no clinical laboratories were using this technology, although individual private veterinary practices sometimes use it. The use of IFM and copro-antigen detection kits in diagnosis is more common elsewhere, particularly in the USA. Two main types of antigen detection system are employed in the kits: enzyme immunoassay assay (EIA) and immunochromatography. The EIA kits are available in multiwell or microplate format, making them an attractive option for laboratories testing large numbers of samples. Indeed, this is the justification for the transfer from microscopy in clinical diagnostics in the UK: laboratories receiving

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large numbers of samples are finding that, particularly if they batch the tests to be performed three times each week, the cost differential with conventional microscopy is reduced. Immunochromatographic tests are available in capsule format, including the dip-stick style for individual sample testing, and offer rapid results within 1–2 hours. Parasites are often shed intermittently and multiple stool samples should be submitted for examination. Traditionally, diagnosis of Giardia in stool, duodenal or jejunal aspirate has been performed by examination of wet preparations (before and after concentration by the formol-ether method) of cysts and trophozoites. Observation is further improved by Lugol’s iodine staining, and examination of a stained permanent smear is also desirable. However, it is doubtful whether sufficient multiple stool samples are submitted for many patients. Vegetative stages in unpreserved stools often degrade, compromising the microscopic diagnosis, but these are still detectable in copro-antigen tests. It has been reported that testing a single stool for copro-antigens by EIA may be an acceptable alternative, since this offers sensitivity approaching that of two sequential stools examined by microscopy (Mank et al., 1997). Improved diagnosis is further enabled by the ‘Triple Faeces Test’ adopted in the Netherlands (van Gool et al., 2003). EIA and immunochromatographic kits are available for individual target pathogens or in combination for Cryptosporidium and Giardia or Cryptosporidium, Giardia and Entamoeba histolytica/dispar. Tests for Cryptosporidium and Giardia can be applied to fresh, frozen or formalin-preserved stools, while those incorporating Entamoeba histolytica/dispar require fresh, unpreserved stools. Sample preparation for antigen detection kits (in contrast to oocyst detection) should not include concentration, since antigens may be lost in the process.

Test Performance Monitoring Should Include Positive and Negative Controls Confirmation of positive test results should always be undertaken, which for microscopy includes accurate measurement of oocysts and observation of morphology. Confirmation of positive results from test kits is important because false positive results have been reported, sometimes driven by high numbers of other microorganisms in the sample. The confirmatory test for diagnostic samples should be of high diagnostic specificity. Most of the test kits were developed using Cryptosporidium parvum: all are useful in detecting Cryptosporidium hominis, but their affinity for other species/genotypes is subject to evaluation.

Test Choice, Evaluation and Quality Assurance The drivers for test choice are theoretically numerous and include cost, simplicity, ease of interpretation, equipment required, personnel/level of training and experience, number of tests ordered, speed, sensitivity, specificity, and the value of incidental findings in non-specific tests. The key attributes of routine test

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options are shown in Table 13.1. However, the main driver is often cost, particularly in publicly funded laboratories. The consumables cost of a modified ZiehlNeelsen test is less than US$1 per sample compared with over US$5 for IFM. However, increasingly, the number of tests ordered is becoming more important as there is a current trend for centralization of laboratory services and there are fewer but larger laboratories testing larger numbers of samples. Here, batch tests such as EIA become more cost-effective when staff costs are taken into account, particularly when the cost of a skilled microscopist’s time is included in the equation. The analytical sensitivity of acid-fast or fluorescence staining of unconcentrated stools is in excess of 104 oocysts per gram (opg), depending on the consistency of the sample, with liquid stools providing greater sensitivity (Anusz et al., 1990; Weber et al., 1991). EIA and immunohistochemical kits offer comparable sensitivity. Improved sensitivity is provided by IFM, but is still in excess of 103 opg. These tests all limit the volume of sample tested. This can be increased to over 2 g by the use of immunomagnetic separation (Robinson et al., 2008) but the cost of this is not justified in routine diagnosis. However, it has a place as a specialist test (see below). The evaluation of new tests should take all the above factors into account and it must also be made clear what is meant by sensitivity and specificity. This is because there are two meanings. One refers to diagnostic sensitivity and specificity and the other to the analytical sensitivity and specificity. Diagnostic parameters are important in studies investigating disease, enabling calculation of positive and negative predictive values for diagnostic tests. The relevant diagnostic definitions are: ●

●

Diagnostic sensitivity = the proportion of subjects with the disease who have a positive test result. Diagnostic specificity = the proportion of subjects with the disease who have a negative test result.

For calculation of these values, there clearly needs to be a clinical definition of disease. However, for Cryptosporidium and Giardia this does not exist in the absence of laboratory confirmation. This presents something of a problem in the above context, which would be only partially overcome by incorporating ‘gold standard’ laboratory confirmation in the definition. Unfortunately, these are not defined either. Nominal definitions may be chosen with justification for individual studies, and Bayesian techniques have also been applied to overcome this problem (Geurden et al., 2006). If a nominal analytical ‘gold standard’ is chosen for comparison of analytical tests, IFM is usually selected for Cryptosporidium and microscopy (wet preparation and stained smear) or IFM for Giardia. Results may be compared using the kappa statistic (k) or McNamar’s test without the necessity to define a gold standard. In studies investigating carriage rates, the analytical sensitivity and specificity are important and are defined thus: ●

Analytical sensitivity = the smallest detectable amount of the analyte in question.

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●

Analytical specificity = the degree to which analytes, other than that in question, react in a test.

Specialist and Reference Tests Routine local primary diagnostic services are often limited to testing stool samples for Cryptosporidium and Giardia, biopsy samples where histological services are available for Cryptosporidium, and duodenal aspirate or string test material for Giardia. Further specialist diagnostic tests may be provided for patients with appropriate indications, or where oocyst numbers are low, either locally or in reference laboratories. The UK Cryptosporidium Reference Unit algorithm recommends specialist testing for patients with idiopathic diarrhoea and/or biliary disease from the following high-risk groups: ● ● ●

HIV with CD4 < 200. Acute leukaemic/lymphoma/BMT. Primary T cell immune deficiency.

Sensitive testing is also available for patients with solid organ transplants or taking T cell immunosuppressants. In addition to stool, appropriate samples types depend on symptoms. For those with cholangitis or biliary disease, stool and bile from ERCP are appropriate and liver biopsy may also be tested. For those with unexplained chest symptoms, sputum or bronchioalveolar lavage may be tested, and for those with unexplained sinusitis, antral washout is appropriate. Test methods are IFM and PCR, which show fair agreement. Improved analytical sensitivity can be provided, where required, by use of immunomagnetic separation (Robinson et al., 2008) or PCR (Webster et al., 1996). Detection using PCR has an emerging place in clinical diagnostics, and is particularly cost-effective when performed as a multiplexed test for two or more pathogens. This approach is being introduced in the Netherlands (L. Kortbeck, personal communication) but the exact choice of which pathogens to co-test for needs careful consideration. None of the routinely used diagnostic tests identifies Cryptosporidium genotypes or Giardia assemblages. Genotyping isolates is currently a specialist test, and provides valuable epidemiological data to measure changes, improve risk assessment, and remove epidemiological noise by separating genotypes in the analysis (Chalmers and Elwin, 2000). Typing can be undertaken from faecal samples or from material fixed on microscope slides in routine diagnostic processing (Amar et al., 2002). It is clear from continuing long-term enhanced surveillance at the UK Cryptosporidium Reference Unit since 2000, involving the testing of over 16,000 isolates, that Cryptosporidium parvum and Cryptosporidium hominis have different epidemiologies (Anonymous, 2002; Nichols et al., 2006) and risk factors (Hunter et al., 2004). Detailed analysis of changes in epidemiology of Cryptosporidium infections is enhanced by genotyping data (Smerdon et al., 2003; Sopwith et al., 2005).

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Reporting and Disease Surveillance While testing for Cryptosporidium is generally voluntary and subject to individual laboratory testing algorithms, reporting for surveillance purposes depends on the notifiable status of the diagnosis in each country. In the UK, Cryptosporidium is not statutorily notifiable unless it is suspected to be foodborne (the definition of which includes waterborne) disease, and reporting is voluntary. Despite this, in 2006 over 80% of laboratories reported cases, indicating the recognized importance of testing and surveillance in the UK. In Ireland, Germany and Sweden, for example, Cryptosporidium is notifiable and reporting compulsory. However, testing is voluntary. Despite advances in the biomedical sciences leading to developments in laboratory diagnostics, examination (usually of faeces) by microscopy remains the keystone of laboratory diagnosis of cryptosporidiosis and giardiasis.

References Amar, C.F.L., Chalmers, R.M., Elwin, K., Tynan, P. and McLauchlin, J. (2002) Blinded evaluation of Cryptosporidium genotyping from DNA recovered from stained smear. Letters in Applied Microbiology 35, 486–488. Anonymous (2002) The Development of a National Collection for Oocysts of Cryptosporidium. Final Report to DEFRA: Drinking Water Inspectorate. Foundation for Water Research, Marlow, Bucks, UK. Available at: http://www.fwr.org/ Anusz, K.Z., Mason, P.H., Riggs, M.W. and Perryman, L.E. (1990) Detection of Cryptosporidium parvum oocysts in bovine feces by monoclonal antibody capture enzyme-linked immunosorbent assay. Journal of Clinical Microbiology 28, 2770– 2774. Arrowood, M.J. (1997) Diagnosis of Cryptosporidium and cryptosporidiosis. In: Fayer, R. (ed.) Cryptosporidium and Cryptosporidiosis. CRC Press, Boca Raton, FL, pp. 43–46. Casemore, D.P. (1991) ACP Broadsheet 128, June 1991: Laboratory methods for diagnosing cryptosporidiosis. Journal of Clinical Pathology 44, 445–451. Casemore, D.P. (1992) Cryptosporidium and Giardia. In: Caul, E.O. (ed.) Immunofluorescent Antigen Detection Techniques in Diagnostic Microbiology. PHLS, London. Casemore, D.P. and Roberts, C. (1993) Guidelines for screening for Cryptosporidium in stools: Report of a Joint Working Group. Journal of Clinical Pathology 46, 2–4. Casemore, D.P., Sands, R.L. and Curry, A. (1985) Cryptosporidium species a “new” human pathogen. Journal of Clinical Pathology 38, 1321–1336. Chalmers, R. and Elwin, K. (2000) Implications and importance of genotyping cryptosporidium. Communicable Disease and Public Health 3, 155–158. Chalmers, R.M., Hughes, S., Thomas, A.L., Woodhouse, S., Thomas, P.D. and Hunter P. (2002) Laboratory ascertainment of Cryptosporidium and local authority public health policies for the investigation of sporadic cases of cryptosporidiosis in two regions of the United Kingdom. Communicable Disease and Public Health 5, 114–118. Farthing, M.J.G. (2000) Clinical aspects of human cryptosporidiosis. In: Petry, F. (ed.) Cryptosporidiosis and Microsporidiosis. Contrib Microbiolo, Karger, Basel, Switzerland, pp. 50–74. Filice, F.P. (1952) Studies on the cytology and life history of a Giardia from the laboratory rat. University of California Publications in Zoology 57, 53–146.

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R.M. Chalmers Garcia, L S. (2001) Diagnostic Medical Parasitology, 4th edn. American Society for Microbiology, Washington, DC. Geurden, T., Berkvens, D., Geldhof, P., Vercruysse, J. and Claerebout, E. (2006) A Bayesian approach for the evaluation of six diagnostic assays and the estimation of Cryptosporidium prevalence in dairy calves. Veterinary Research 37, 671–682. Hunter, P.R. and Nichols, G. (2002) Epidemiology and clinical features of Cryptosporidium infection in immunocompromised patients. Clinical Microbiology Reviews 15, 145– 154. 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. Emerging Infectious Diseases 10, 1241– 1249. Kofoid, C.A. (1920) A critical review of the nomenclature of human intestinal flagellates Cercomonas, Chilomastix, Trichomonas, Tetratrcihomonas, and Giardia. University of California Publications in Zoology 20, 145–168. Kofoid, C.A. and Christensen, E.B. (1915) On binary and multiple fission in Giardia muris (Grassi). University of California Publications in Zoology 16, 30–54. Lambl, W. (1859) Mikroskopische untersuchungen der Darmexcrete. Vierteljahrsschr Prakst Heikunde 61, 1–58. Mank, T.G., Zaat, J.O.M., Deelder, A.M., van Eijk, J.T.M. and Polderman, A.M. (1997) Sensitivity of microscopy versus enzyme immunoassay in the laboratory diagnosis of giardiasis. European Journal of Clinical Microbiology and Infectious Diseases 16, 615–619. Nichols, G. and Thom, B.T. (1984) Screening for Cryptosporidium in stools. Lancet 1, 735. 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 Inspectorate Contract Number DWI 70/2/201. Drinking Water Inspectorate, London. Robinson, G., Watkins, J. and Chalmers, R.M. (2008) Evaluation of a modified semiautomated immunomagnetic separation technique for the detection of Cryptosporidium oocysts in human faeces. Journal of Microbiological Methods 75, 139–141. Smerdon, W.J., Nichols, T., Chalmers, R.M., Heine, H. and Reacher, M.H. (2003) Decrease in human cryptosporidiosis coincident with the Foot and Mouth Disease epidemic: a measure of the livestock attributable fraction in England and Wales? Emerging Infectious Diseases 9, 22–28. Sopwith, W., Osborn, K., Chalmers, R. and Regan, M. (2005) The changing epidemiology of cryptosporidiosis in north west England. Epidemiology and Infection 133, 785–793. Tyzzer, E.E. (1907) A sporozoan found in the peptic glands of the common mouse. Proceedings of the Society for Experimental Biology and Medicine 5, 12–13. Tyzzer, E.E. (1910) An extracellular coccidium, Cryptosporidium muris (gen. et sp. nov.) of the gastric glands of the common mouse. Journal of Medical Research 23, 487–509. Tyzzer, E.E. (1912) Cryptosporidium parvum (sp. nov.), a coccidium found in the small intestine of the common mouse. Archiv fur Protistenkunde 26, 394–412. Tyzzer, E.E. (1929) Coccidiosis in gallinaceous birds. American Journal of Hygiene 10, 269–383. van Gool, T., Weijts, R., Lommerse, E. and Mank, T.G. (2003) Triple faeces test: an effective tool for detection of intestinal parasites in routine clinical practice. European Journal of Clinical Microbiology and Infectious Diseases 22, 284–290.

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Weber, R., Bryan, R.T., Bishop, H.S., Walquist, S.P., Sullivan, J.J. and Juranek, D.D. (1991) Threshold of detection of Cryptosporidium oocysts in human stool specimens: evidence for low sensitivity of current methods. Journal of Clinical Microbiology 29, 1323–1327. Webster, K.A., Smith, H.V., Giles, M., Dawson L. and Robertson, L.J. (1996) Detection of Cryptosporidium parvum oocysts in faeces: comparison of conventional coproscopical methods and the polymerase chain reaction. Veterinary Parasitology 61, 5–13.

14

Control of Cryptosporidium and Giardia in Surface Water by Disinfection

T.M. HARGY1, J.L. CLANCY1 AND L.P. LANDRY2 1Clancy

Environmental Consultants, Saint Albans, VT, USA; 2Greater Vancouver Regional District, Burnaby, Canada

Abstract The Greater Vancouver Regional District (GVRD) serves the Greater Vancouver, BC, area, delivering water to 18 Lower Mainland municipalities, which in turn deliver water to approximately 2.1 million people. Water is collected from three mountainous watersheds: Capilano, Coquitlam and Seymour. The GVRD adopted its Drinking Water Management Plan (DWMP) in August 2005 and its strategy is to use a risk-management multi-barrier approach from source to tap. The results of this project have confirmed that the GVRD has selected an effective multi-barrier approach for protecting the health of consumers of unfiltered drinking water. However, caution is recommended if any circumstance arise such that turbidity spikes due to some cause that enriches the organic, rather than the inorganic, fraction of the suspended particles. An algal bloom or terrestrial upset, such as the immediate fallout of a forest fire, might have an effect not tested in this study. The protected nature of the watershed makes a nutrient influx fuelling an algal bloom an unlikely scenario, but a natural or human-induced forest fire is possible. Ozone was shown to be a very effective pretreatment of Coquitlam water for UV, regardless of turbidity. The UV transmittance (UVT) of Coquitlam water, and thus its UV treatability, was uniformly improved by ozonation.

Introduction The Greater Vancouver Regional District (GVRD) serves the Greater Vancouver, BC, area, delivering water to 18 Lower Mainland municipalities, which in turn deliver water to approximately 2.1 million people. Water is collected from three mountainous watersheds: Capilano, Coquitlam and Seymour. The GVRD adopted its Drinking Water Management Plan (DWMP) in August 2005 and its strategy is to use a risk-management multi-barrier approach from source to tap. The Coquitlam watershed is closed to the public, industry and agriculture to minimize potential contamination of the source water. The Coquitlam source is currently using ozone for primary disinfection and chlorine for secondary disinfection. The GVRD plans to install UV for Cryptosporidium control downstream of the 158

© CAB International 2009. Giardia and Cryptosporidium: From Molecules to Disease (eds G. Ortega-Pierres et al.)

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ozone treatment for Giardia and viruses. While Coquitlam source water turbidity is low and filtration can be avoided in accordance with the Guidelines for Canadian Water Quality, there may be infrequent, but possible, events likely to cause high source-water turbidity. This study was undertaken to demonstrate the degree to which an elevated turbidity event would diminish the effectiveness of the disinfection system. Specifically, the question asked was whether elevated turbidity at the Coquitlam source could directly impair the effectiveness of either the UV disinfection of Cryptosporidium or the ozone disinfection of Giardia and viruses. A second concern was whether elevated turbidity might minimize the advantage gained when UV is applied downstream of ozone. In this study, bulk water samples and concentrated suspended material were collected and used to quantify and characterize the microbiological and physical characteristics of Coquitlam source water. Further efforts were directed toward re-suspending subsamples of the collected concentrated material into reactor volumes of the bulk water, and evaluating the efficacy of ozone and UV at disinfecting natural or seeded microorganisms across a range of turbidities experimentally prepared from native Coquitlam particles. Specifically, we examined the disinfection by ozone and UV of seeded coliphage MS2, Cryptosporidium oocysts and Giardia cysts. Finally, we investigated the effect of ozone on UV absorbance to determine whether the oxidant would continue to improve the UV treatability of Coquitlam water under high turbidity conditions, to the same degree as it has been shown to do at the lower turbidity levels generally present.

Enumeration of Indigenous Free- and Particle-associated Microorganisms in Coquitlam Water A combination of methods was used to assess the microbiological quality of the Coquitlam source water that supplies Greater Vancouver’s water needs. Knowing the types and concentrations of microorganisms present in a source water is important in assessing the type(s) of treatment necessary to comply with drinking water regulations or guidelines and to protect public health. Organisms of interest include pathogens such as Giardia and Cryptosporidium, relatively resistant organisms such as bacterial spores, algae – which are often the most abundant organisms in a waterbody, and macroorganisms that include a variety of invertebrates that are not pathogenic but contribute to the total biomass. Macro- and microorganisms are charged particles and tend to become associated with other particles, forming aggregates. To characterize the water quality, both the levels and types of biota present under various conditions were examined. The methods used to measure the microbiological quality of the Coquitlam water supply included direct microscopic observation using a technique known as microscopic particulate analysis or MPA; USEPA method 1623 (USEPA, 2005) for detection of Giardia cysts and Cryptosporidium oocysts; heterotrophic plate count (HPC), a measure of general microbial quality; total aerobic spores (TAS), which measures the population of aerobic resistant spore-forming bacteria; and male-specific and somatic coliphages, bacterial viruses that infect E. coli.

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Sample collection Samples for subsequent testing and analyses were collected by GVRD from the Coquitlam source immediately downstream of the utility intake at a sample port.

Giardia/Cryptosporidium and MPA Analyses Filter processing For each high-volume sample, a spiral wound filter and two Envirochek HV capsule filters were collected. The spiral wound filter was processed as described in the MPA consensus method (USEPA, 1992) and the HV filters were processed as described in USEPA method 1623 (USEPA, 2005), with the exception that, for the purpose of characterizing the microbial population of the captured particulate material, each filter was processed using an aseptic technique. MPA analysis A portion equivalent to 0.7 l of each sample collected was examined using bright field microscopy. Giardia/Cryptosporidium enumeration Low-volume samples were collected on Envirochek HV filter capsules. The retained particles were eluted from the filter as described above, with the final concentrate being resuspended to 10–20 ml. A portion equivalent to 10–15 l of the original sample volume was transferred for Cryptosporidium and Giardia analysis by immunomagnetic separation (IMS). Recovered organisms were enumerated using epifluorescence microscopy. Pellet preparation for microbial analysis Packed pellet material was gently resuspended and an aliquot (1% of this original suspension) was diluted in sterile PBW and mixed. This sample was split into three subsamples, one for each of the three microbial assays (HPC, TAS and coliphages). Each subsample was split in half: one half was analysed directly, and the other half was analysed following a disassociation procedure involving a series of vortexing and sonication steps. Enumeration of heterotrophic bacteria Heterotrophic bacteria were enumerated by the spread plate technique (Standard Method 9215C) on R2A agar to measure HPC. Plates were incubated inverted

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for 4–7 days at room temperature (25°C). Colony forming units (CFU) from dilutions were recorded and the concentration of HPC was calculated as CFU/ml.

Enumeration of total aerobic spores TAS were enumerated by the spread plate technique (Standard Method 9215C) on TAS medium. Following pasteurization, samples were subjected to serial tenfold dilution in PBW (if necessary). An appropriate volume of sample water was aseptically spread on TAS medium using a sterile spreader and allowed to absorb completely before incubating. Plates were incubated inverted for 24 ± 2 h at 35 ± 0.5°C. Counts were recorded and the TAS concentration calculated as CFU/ml.

Enumeration of male-specific and somatic coliphages Male-specific and somatic coliphage concentrations were determined by the double-agar-layer plaque assay (Standard Methods 9224B and 9224C) using host bacteria Escherichia coli HS(pFamp)R (ATCC No. 700891). Viral plaque counts from dilutions in the countable range were tallied and reported as plaque forming units per ml (PFU/ml).

Results Giardia and Cryptosporidium Data for pathogenic protozoa are shown in Table 14.1. No fluorescing objects resembling Giardia cysts or Cryptosporidium oocysts were observed in either of the samples analysed.

Microscopic particulate analysis Two MPA samples were collected and analysed using the USEPA consensus method (USEPA, 1992). Results are shown in Table 14.2. The increase in turbidity from Coquitlam 1 to Coquitlam 2 led to a large increase in packed pellet volume; the Coquitlam 1 sample had a packed pellet Table 14.1.

Giardia and Cryptosporidium sampling data.

Sampling round Sample matrix Coquitlam 1 Coquitlam 2

Envirochek HV capsule Envirochek HV capsule

Giardia cysts Cryptosporidium oocysts <1/10 l <1/15 l

<1/10 l <1/15 l

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Microscopic particulate analysis data.

Sample date Volume sampled (l) Start/end turbidity (NTU) Packed pellet volume (ml)

Coquitlam 1

Coquitlam 2

11/7/2006 400

11/16/2006 124

1.5/2.0

6.2/5.3

0.5

4

(No. per 100 l)

(No. per 100 l)

Algae

5.7 × 106

7.0 × 106

Amoebae

1.1 × 104

6.2 × 103

Crustaceans

MPA category

1.4 ×

102

ND

Diatoms

3.1 ×

104

3.4 × 104

Insects/parts

2.0 ×

103

2.8 × 102

Nematodes/eggs

2.4 × 103

7.1 × 102

Pollen

5.7 × 102

3.4 × 103

ND

ND

Free-living protozoa Rotifers/eggs Spores Vegetative debris

8.5 ×

102

4.3 × 102

ND

2.3 × 103

1.4 × 102

1.8 × 103

ND: none detected.

volume of 0.5 ml/400 l (0.1 ml/100 l) while Coquitlam 2 had a 4 ml pellet from a 124 l sample volume (3.3 ml/100 l). Coquitlam 1 contained biological particles typical of pristine lake water, with low levels of algae and diatoms. Inorganic debris in the sample was typical of lake water, with bits of diatom shells adding to the ‘inorganic’ or solids portion of the sample. The amount of inorganic debris present was small (<15 µm), colourless and fairly sparse. When a portion of the sample was concentrated by centrifugation, a layer of algae was observed on top of the inorganic portion of the sample. This observation is unusual; however, if algae are present that form gas vacuoles or are contained in colonial mucilage, this phenomenon can occur. The algae layer comprised approximately 10% of the volume of the pellet material present in this sample. Coquitlam 2 contained a much larger number of inorganic particles. The concentrations and types of biological particles in this sample were similar to the Coquitlam 1 sample. The biological portion of the sample comprised <0.1% of the particle concentration in this sample. While similar concentrations of algae and diatoms were noted in both MPA samples examined, the amounts of inorganic debris increased significantly in the higher-turbidity sample. The number of inorganic particles increased by a log, indicating that the increase in turbidity did not result in an increased load of biological particles, but rather in inorganic particles due to runoff. Concentrations of pollen and vegetative debris were

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greater in Coquitlam 2; while these are biological in nature, they are also likely to be increased due to runoff during the rainfall event. The photomicrographs in Fig. 14.1 show the difference observed in the two samples. The Coquitlam 1 sample (Photos 1A–1D) had much less inorganic debris than did Coquitlam 2 (Photos 2A–2D).

Heterotrophic bacteria HPC data are shown in Table 14.3. The HPC concentrations were enumerated from the 1-l bulk water samples collected aseptically immediately on receipt of the samples, and the counts were similar for both samples (1.5–1.8 × 103/ml). The pre- and post-disaggregation sample data were measured on the packed pellet material and represent hundreds of litres of filtered water concentrate; they do not relate to the bulk water data in the table. The pellet counts show that there was no increase in HPCs after the sample was subjected to vortexing and sonication, suggesting that particle absorption did not result in clumping of HPC bacteria, and thus cause an underestimation of their presence.

Total aerobic spores Table 14.4 shows the TAS data. The spore data show that TAS was very low or not detected in all of the samples tested. Even the packed pellet material, which is hundreds-fold concentrated, showed no spores before or after treatment to disrupt aggregates. Spores are highly resistant to inactivation but their low numbers in this source water would not allow their use as a natural indicator of disinfection.

Male-specific and somatic coliphage Coliphage data are shown in Table 14.5. Like TAS, the phage data showed very low counts in both samples of the bulk water. The packed pellet concentrates were very low and disaggregation did not increase counts. Like TAS, coliphages would not be a useful natural indicator of disinfection efficacy.

Disinfection of MS2, Cryptosporidium and Giardia Project approach In this phase of the project we characterized the dose response of coliphage MS2, Giardia and Cryptosporidium in baseline Coquitlam water to a range of disinfectant doses, with the purpose of identifying the dose necessary for 4-log (MS2) and 3-log (protozoa) inactivation. This was followed by testing the efficacy of that disinfectant dose at inactivating the microorganisms suspended in

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Coquitlam 1A - Minimal inorganic debris; centric diatom, girdle view; Cyanophyta; 200

Coquitlam 1B - Minimal inorganic debris, Pleurotaenium; 200

Coquitlam 1C - Aggregated inorganic debris, 400

Coquitlam 1D – Merismopedia and Chroococcus, 400

Coquitlam 2A - Inorganic debris with amoebae shell, 200

Coquitlam 2B - Pennate diatom and Chroococcus, 400

Coquitlam 2C - Merismopedia, 400

Coquitlam 2D- Inorganic debris, 400

Fig. 14.1. Photomicrographs of particles in Coquitlam waters 1 and 2, showing the differences noted in each sample.

Control of Cryptosporidium and Giardia in Surface Water Table 14.3.

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Heterotrophic plate count data.

Sampling round

Sample matrix

Average CFU/ml

Standard deviation

1 2 1 2 1 2

1 l bulk water 1 l bulk water Pellet pre-disaggregation Pellet pre-disaggregation Pellet post-disaggregation Pellet post-disaggregation

1.8E+03 1.5E+03 6.3E+05 7.2E+06 6.0E+05 7.3E+06

1.2E+02 1.0E+02 7.6E+04 1.2E+05 9.8E+04 1.2E+05

Average CFU/ml

Standard deviation

0.3 0.1 0 0 0 0

0.1 0 0 0 0 0

Table 14.4. Total aerobic spore data. Sampling round

Sample matrix

1 2 1 2 1 2

1 l bulk water 1 l bulk water Pellet pre-disaggregation Pellet pre-disaggregation Pellet post-disaggregation Pellet post-disaggregation

Table 14.5.

Coliphage data.

Sampling round

Sample matrix

1 2 1 2 1 2

1 l bulk water 1 l bulk water Pellet pre-disaggregation Pellet pre-disaggregation Pellet post-disaggregation Pellet post-disaggregation

Male-specific phage

Somatic phage

Average PFU/ml

Average PFU/ml

0.2 <0.3 <0.01 <0.003 <0.01 <0.003

0.2 <0.3 <0.01 <0.003 0.01 <0.003

Coquitlam water that had previously been adjusted to a range of turbidity levels in order to evaluate any degradation of disinfection with increase in turbidity. The turbidity levels selected were from baseline up to 5 NTU in 1-NTU increments, for evaluation of disinfection efficacy across the turbidity range permissible under the Guidelines for Canadian Drinking Water Quality (Health Canada, 2005); also selected were 10, 15 and 20 NTU for evaluation of disinfection in the face of currently potential operating conditions (up to the 99.995% percentile value of 20 NTU) wherein the Coquitlam source could not be shut down during turbidity events. Such a scenario will no longer present itself on completion of the Seymour Capilano filtration plant. Finally, the 50 NTU level was selected as an extraordinary turbidity level at which to observe any impacts to disinfection practices.

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Turbidity adjustment In all disinfection against turbidity tests, the test microbes were mixed with sufficient particulate material to achieve the target turbidity in the reactor volume, and allowed to stand overnight at 4°C to allow contact time for particle association. Sufficient MS2 were added to result in reactor concentrations of 1 × 107 PFU/ ml for UV tests and 2 × 106 PFU/ml for ozone tests. Protozoa were added in quantities producing a total of 1 × 106 microorganisms per reactor volume. At the time of testing, agitation of the microbe/particle mixture for resuspension was kept to a minimum to avoid breaking up any aggregates that may have formed.

UV disinfection methods Collimated beam dose response determination The UV dose delivery process, known as a collimated beam test, was carried out using the methods described by Bolton and Linden (2003), which adjusts exposure time based on UV irradiance, absorbance and distribution. UV dose was defined as the average irradiation so calculated, multiplied by the exposure time. Exposures providing UV doses of 0, 20, 40, 60, 80 and 100 mJ/cm2 for MS2, and 0, 1, 2, 3, 4 and 5 mJ/cm2 for Cryptosporidium were used. A control (0 mJ/ cm2 dose) was run simultaneously with the longest exposure for each microbe, but in the absence of UV. The control sample provided the base count for determination of log inactivation. UV dose selection and determination in the presence of turbidity The same collimated beam apparatus and methods employed for dose response determination were used to evaluate the effects of turbidity on UV disinfection of MS2 and Cryptosporidium, but with the following adjustments. Rather than present a range of exposure times, and thus UV doses, to suspended organisms, the UV dose determined from the above dose response characterization was applied to the test microbes suspended in a range of turbidities from baseline to as high as 50 NTU. As noted above, in addition to inoculating the baseline water with test organisms, Coquitlam particulate material was added to the reactor volumes and left overnight to achieve increments of target turbidity levels. For UV disinfection tests, the direct UV absorbances at 254 nm were measured at each turbidity level to quantify the water correction factors for each exposure time.

Microorganism propagation and assay MS2 was selected because it has been shown to be a conservative surrogate for enteric virus disinfection by UV (Wilson et al., 1992), though it is more sensitive to UV than is adenovirus (Meng and Gerba, 1996). Its UV dose response is well documented (USEPA, 2006). For the present project, MS2 (ATCC No. 15597-B1) was propagated in E. coli Famp, using a liquid culture method. Prior to disinfection challenges, the stock was diluted in sterile PBW to achieve a working stock

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concentration of 1 × 109 PFU/ml. This stock was dispensed to reactor batches in volumes sufficient to achieve 1 × 107 PFU/ml reactor suspensions. MS2 assays Phage concentrations were determined via the double agar layer technique (Adams, 1959). All samples were plated in triplicate. PFU were counted and recorded, and the concentration of MS2 calculated. Disaggregation following UV exposure in elevated NTU In analysing the MS2 surviving UV in elevated turbidity Coquitlam water, dissociation steps were employed to dissociate the phage from the particulate matrix in subsamples of one test set. Cryptosporidium stock Cryptosporidium parvum oocysts (Iowa isolate) were obtained from Waterborne, Inc. (New Orleans, LA). Oocysts were fresh (16 days old post shedding) at the time of testing. Cell culture assay Samples to be analysed for infectious Cryptosporidium oocysts were processed as described by M.S. Johnson (Water Quality Laboratory, Metropolitan Water District of Southern California, 2006, personal communication). UV disinfection results MS2 UV dose response The responses of MS2 seeded into Coquitlam waters at baseline (0.3 NTU) turbidity are presented in Figs 14.2 and 14.3 and indicate that MS2 suspended in MS2 UV dose response curve, Seeded Coquitlam 1 water, baseline conditions (low NTU) 100 UV dose (mJ/cm2)

y = 0.9055x 2 + 17.267x 80 60 40 20 0 0

1

2 3 Log inactivation

4

5

Fig. 14. 2. The UV dose response of MS2 seeded into Coquitlam 1 water at baseline (0.3 NTU) turbidity.

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Coquitlam 1 required 83 mJ/cm2 for 4-log inactivation, while MS2 in the Coquitlam 2 required 92 mJ/cm2 for the same level of disinfection. On introducing particulate matter from the respective samples to achieve elevated turbidity, UV sensitivities at the doses required for 4-log inactivation (83 and 92 mJ/cm2 for Coquitlam 1 and 2, respectively) were found to be as effective as in baseline waters (Figs 14.4 and 14.5). In the latter elevated turbidity test, the sample was disaggregated after UV irradiation to disassociate MS2 from particles. Cryptosporidium disinfection by UV The inactivation of Cryptosporidium oocysts suspended in baseline Coquitlam water is shown in Fig. 14.6. More than 3.6-log inactivation was achieved at 3 mJ/cm2 and more than 5-log inactivation was demonstrated at 4 and 5 mJ/cm2. As 3 mJ/cm2 achieved the GVRD target of 3-log Cryptosporidium inactivation, that UV dose was then applied to seeded reactors with incrementally elevated

MS2 UV dose response curve, Seeded Coquitlam 2 water, baseline conditions (low NTU) 100 UV dose (mJ/cm2)

y = 0.9787x 2 + 19.151x 80 60 40 20 0 0

1

2 3 Log inactivation

4

5

Fig. 14.3. The UV dose response of MS2 seeded into Coquitlam 2 water at baseline (0.3 NTU) turbidity.

UV log inactivation of MS2 vs turbidity Coquitlam sample 1, 83 mJ/cm2 Log inactivation

5.0 4.0 3.0 2.0 1.0 0.0 0

5

10

15 NTU

20

25

(50)

Fig. 14.4. The UV inactivation of MS2 versus turbidity in Coquitlam 1 water.

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UV log inactivation of MS2 vs turbidity Coquitlam sample 2, 92 mJ/cm2 Log inactivation

5.0 4.0 Raw Disaggregated

3.0 2.0 1.0 0.0 0

5

10

15

20

25

(50)

NTU

Fig. 14.5. The UV inactivation of MS2 versus turbidity in Coquitlam 2 water.

UV dose response of Cryptosporidium in baseline Coquitlam water 6

Log inactivation

5 4 3 2 1 0 0

1

2

3

4

5

UV dose (mJ/cm2)

Fig. 14.6. The UV dose response of Cryptosporidium parvum oocysts seeded into Coquitlam baseline water.

turbidity for each water and pellet sample. The results, shown in Table 14.6, indicate that 3-log inactivation of Cryptosporidium by UV was not diminished when oocysts were seeded overnight in Coquitlam particulate matter and subsequently exposed to UV.

Ozone disinfection of MS2 and Giardia Ozone generation and ozone stock production Ozone was produced using a Clearwater Technology CD10/AD corona discharge ozone generator that was modified to accept feed gas from a pressurized source. The feed gas was dry compressed air. The output was delivered to 1 l of 3°C Coquitlam water in an ozone demand-free side arm flask through a glass diffuser

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T.M. Hargy et al. Table 14.6. The UV inactivation of Cryptosporidium with elevated Coquitlam turbidity at 3 mJ/cm2.

Turbidity (NTU) 0 5 10 20 50

Log inactivation in Coquitlam sample 1

Log inactivation in Coquitlam sample 2

>5 >5 >5 >5 >5

>5 >5 >5 >5 >5

inserted through a single-hole stopper. Off-gas was vented from the side arm port and exhausted from the building. Ozone residual measurement All ozone residual analyses were made using a variation of Standard Method 4500-O3 B. To compute the integrated products of disinfectant residual and contact time, ozone residuals recorded over the course of the experiment were multiplied by the contact time interval between each residual measurement. Ozone reactors Batch reactors were prepared by seeding test water in 100 ml ozone demandfree beakers containing stir bars and situated in a water bath (at 3–4°C) over a stir plate. After determining the ozone concentration of the ozone stock, sufficient stock to achieve the target applied ozone or initial ozone residual was pipetted at time zero to the reactors, previously seeded and containing the appropriate volume of test water such that addition of ozone stock would result in a total volume of 50 ml. Ozone reactivity was quenched by adding sodium bisulphite. The effectiveness of the quenching reaction was confirmed by addition of the bisulphite solution to a test reactor containing 3 mg/l ozone and measuring a 0 mg/l ozone residual. The quenching agent was also added to the process control to which no ozone stock had been added. Giardia muris preparation Giardia muris was obtained from the Meyer Laboratory, Oregon Health Sciences, Portland, OR. The purified cysts were stored at 3–5°C in antibiotics for up to 2 weeks post shedding. Sample processing and excystation procedure Samples were received in the laboratory in 40–45 ml volumes and subjected to an excystation procedure as described by Hoff et al. (1985). Cysts were scored as excysted (empty cysts), partially excysted (trophozoite emerging from cyst), and non-excysted (intact cyst containing trophozoite). Cysts which were scored as excysted and partially excysted were considered viable, and those that did not excyst were scored as non-viable.

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Ozone disinfection results MS2 Figure 14.7 shows the inactivation kinetics of MS2 seeded in baseline Coquitlam water and subjected to ozone CT values (concentration of ozone multiplied by contact time with virus) from 0.01 to 0.3 mg min/l. From the resulting curve, the 4-log inactivation benchmark would be achieved by a CT of 0.26 mg min/l. This 0.26 CT level and a slightly higher CT (0.35) were then applied to 50 ml reactor volumes adjusted across the range of turbidity including 0, 2, 5, 10 and 50 NTU with Coquitlam particulate material that had been seeded overnight with MS2 stock (2 × 106 PFU/ml of reactor volume). In all tests, inactivation proceeded to the point where no surviving phages were detected in the highest inoculum volume we tested, demonstrating inactivations of greater than 5.5-log. A lone exception was that a single plaque, indicating one surviving phage, was detected in the lower CT test at 10 NTU, resulting in a reportable log inactivation of 5.5, rather than greater than that level. Ozone disinfection of Giardia Ozone disinfection of Giardia cysts was carried out for both waters under baseline and elevated turbidity conditions. The ozone dose responses of Giardia are shown in Fig. 14.8; a CT of 3.3 mg min/l achieved greater than 3-log inactivation in both waters. That CT level was then targeted to the elevated turbidity reactors. The results, shown in Table 14.7 indicate that for both Coquitlam waters, more than 3-log inactivation was achieved by a CT of approximately 3 (range: 2.7– 3.2) in turbidities of baseline and 5–20 NTU. Evaluation of increasing turbidity effect on ozone/UV synergy The UV transmittance (UVT) of baseline Coquitlam waters (254 nm) and waters adjusted to 5–50 NTU were measured across 1 cm at Clancy Environmental MS2 ozone dose response curve, seeded baseline Coquitlam water

Ozone CT (mg min/L)

0.4 0.3 0.2 0.1 0 0

1

2 3 Log inactivation

4

5

Fig. 14.7. The inactivation kinetics of MS2 seeded into baseline Coquitlam water and subjected to ozone CTs from 0.01 to 0.3 mg min/l.

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T.M. Hargy et al. Log inactivation of Giardiamuris cysts vs ozone CT in baseline Coquitlam waters 1 and 2

Log inactivation

4 3 2 1 0 0

0.5

1

1.5 2 CT (mg min/l)

2.5

3

3.5

Fig. 14.8. The ozone dose responses of Giardia muris cysts seeded into baseline Coquitlam waters 1 and 2. Table 14.7. Ozone disinfection of Giardia muris cysts seeded into Coquitlam water adjusted to a range of turbidities at 3.5°C. Coquitlam sample number

Turbidity (NTU)

CT (mg min/L)

Log inactivation

1 1 1 1 2 2 2 2

Baseline (0.3) 5 10 20 Baseline (0.3) 5 10 20

3.1 2.8 2.7 3.0 2.9 3.2 2.9 3.2

>3.0 >3.0 >3.0 >3.0 >3.0 >3.0 >3.0 >3.0

Consultants (CEC) using direct spectrophotometry (Spectronic Genesys 10UV). Split samples of these waters were ozonated with an applied ozone dose of 2.4 mg/l, and UV absorbances were measured again. Pre- and post-ozone split samples of each of these turbidity levels were shipped chilled overnight to the Department of Civil and Environmental Engineering at Duke University and direct transmittance was again measured using a Cary 300 spectrophotometer. Finally, an integrated sphere device was incorporated to allow measurement of UV absorbance without interference from light scattering by particles. The results of these analyses are shown graphically in Figs 14.9 to 14.12. Figures 14.9 and 14.10 present the pre- and post-ozonated direct UVT data for Coquitlam samples 1 and 2, as measured by CEC and Duke University, and indicate the change in direct UVT as a result of ozonation across the range of turbidities tested. Generally, pre-ozone baseline UVT values were 86%, and decreased with turbidity to as low as 52% at 50 NTU. These figures indicate a significant improvement in UVT post-ozonation in both waters across the turbidity range, with UVT improving in the baseline water by as much as 8 percentage points to 94%.

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Coquitlam 1 direct UV 254% transmittance, pre- and post-ozone, two laboratories 100

CEC DUKE

UV 254%T 1 cm

90 80

Direct %T, Post-ozone

70 Direct %T, Pre-ozone

60 50 0

10

20

30

40

50

Turbidity (NTU)

Fig. 14.9. The pre- and post-ozonated direct UVT data for Coquitlam sample 1, as measured by Clancy Environmental Consultants (CEC) (direct spectrophotometry; Spectronic Genesys 10UV) and Duke University (Cary 300 spectrophotometer). Coquitlam 2 direct UV 254% transmittance, pre- and post-ozone, two laboratories

UV 254%T, 1 cm

100

CEC DUKE

90

Direct %T, Post-ozone 80 70 Direct %T, Pre-ozone 60 50 0

10

20 30 Turbidity (NTU)

40

50

Fig. 14.10. The pre- and post-ozonated direct UVT data for Coquitlam sample 2, as measured by Clancy Environmental Consultants (CEC) (direct spectrophotometry; Spectronic Genesys 10UV) and Duke University (Cary 300 spectrophotometer).

Figures 14.11 and 14.12 compare the integrated sphere (true) pre- and postozone UVT of Coquitlam 1 and 2 waters, respectively. As was noted with direct UVT measurements, ozonation was a beneficial pretreatment for UV. The advantage widens with turbidity in Coquitlam 1, but narrows in Coquitlam 2.

Discussion UV Disinfection of MS2 and Cryptosporidium The sensitivity of MS2 to UV in baseline Coquitlam water was in line with the findings of the numerous studies evaluated in USEPA’s Ultraviolet Disinfection

174

T.M. Hargy et al. Coquitlam 1 integrated sphere (true) UV 254% transmittance, pre- vs post-ozone

UV 254%T, 1 cm

100 90

True %T, Post-ozone

80 70 True %T, Pre-ozone 60 50 0

10

20

30

40

50

Turbidity (NTU)

Fig. 14.11. The integrated sphere (true) pre- and post-ozone UVT of Coquitlam 1 water. Coquitlam 2 integrated sphere (true) UV 254% transmittance, pre- vs post-ozone

UV 254%T, 1 cm

100 True %T, Post-ozone

90 80 True %T, Pre-ozone

70 60 50 0

10

20 30 Turbidity (NTU)

40

50

Fig. 14.12. The integrated sphere (true) pre- and post-ozone UVT of Coquitlam 2 water.

Guidance Manual (UVDGM) (2006). As shown in Table 14.8, all data points generated with baseline Coquitlam water fell well within the recommended limits of the UVDGM. A dose of 83 mJ/cm2 was found to achieve 4-log inactivation of MS2 seeded into the first Coquitlam sample, while 92 mJ/cm2 was required in the second test water. The difference between the curves from the two sample dates is within the range of variability typically seen between collimated beam curves, and is not thought to be indicative of any difference in treatability of the two waters. This interpretation is supported by the results of the test wherein the 4-log inactivation doses (83 and 92 mJ/cm2) were applied to elevated turbidity reactor volumes. In the case of Coquitlam 1, 4-log inactivation was achieved fairly uniformly (range: 3.8–4.1 log) in waters of 0 to 50 NTU. In the second water test, the application of 92 mJ/cm2 uniformly achieved well over 4-log inactivation (range 4.2–4.7).

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Table 14.8. Comparison of MS 2 UV dose response in Coquitlam waters with UVDGM recommendations (USEPA, 2006). UV dose (mJ/cm2) 20 40 60 80 100

Coquitlam 1

Coquitlam 2

UVDGM lower bounds

UVDGM upper bounds

1.2 2.1 3.0 3.7 4.7

1.1 2.0 2.7 3.4 4.3

0.9 1.7 2.4 3.0 3.5

1.5 2.8 4.1 5.2 6.2

This suggests that the 92 mJ/cm2 value determined in the baseline dose response curve for Coquitlam water 2 was high. This is further confirmed by the fact that these higher inactivations achieved in elevated Coquitlam 2 turbidity would be expected by a dose of about 89–100 mJ/cm2 from the first Coquitlam baseline curve. The inactivation data shown in Figs 14.4 and 14.5 indicate that there was no strong negative impact to the inactivation of coliphage MS2 seeded overnight in Coquitlam water at elevated turbidities. That slight shielding did occur is indicated by the results of UV-exposed samples that were put through a dissociation regimen prior to assay. As this process released viable MS2 from the treated matrix, increased counts per ml were realized, decreasing the log inactivation achieved. This exercise, performed on Coquitlam 2 tests from 10 to 50 NTU, demonstrated actual inactivation to be 0.1-log less in all instances. A remaining uncertainty is whether naturally occurring, naturally shielded viral pathogens might pass unaffected through a UV reactor. The low concentrations of indigenous phage and the dominance of mineral, inorganic particles would suggest this is not likely to be a concern for a UV system treating Coquitlam water. Cryptosporidium This study has corroborated the findings of Clancy et al. (2004) and Shin et al. (2001) that approximately 1 mJ/cm2 is necessary for 1-log inactivation of Cryptosporidium. These two references, along with others, form the basis for the USEPA’s inactivation table in UVDGM LT2 (USEPA, 2006), which takes data variability into account and applies safety factors in granting 1-, 2-, 3- and 4-log inactivation credits for validated UV doses of 2.5, 5.8, 12 and 22 mJ/cm2, respectively. Ozone disinfection of MS2 and Giardia The sensitivity to ozone noted here for MS2 masked any impact by Coquitlam particles, as the steepness of the inactivation kinetics exceeded the experimental resolution of CT determination. The application of a CT only 0.1 mg min/l greater than that found to be necessary for 4-log inactivation resulted in at least 5.5-log inactivation across all turbidities.

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The ozone CT table in the Canadian Guidelines for Drinking Water Quality (Health Canada, 2005) requires CTs of 1.9 and 2.9 mg min/l at 5°C and 1°C, respectively, for 3-log ozone inactivation of Giardia cysts. Assuming a linear relationship between temperature and CT, the CT for 3-log inactivation at 3.5°C would be 2.5 mg min/l, which is roughly the effective CT determined here for 3-log inactivation in baseline Coquitlam water. The results of testing in Coquitlam waters 1 and 2 indicate that ozone is effective in Coquitlam waters at 3.5°C and across a range of turbidities. That no apparent impact of turbidity on ozone efficacy could be discerned from the results of this study is supported by the findings of Dow et al. (2006) in their investigation of ozone disinfection of relatively ozone-resistant B. subtilis spores under varying water quality conditions, including turbidity.

Effect of turbidity on ozone and UV UVT measurements indicate that ozonation of Coquitlam water with an applied dose of 2.4 mg/l will be a highly beneficial pretreatment for UV disinfection. Comparisons of direct UVT values measured in pre- and post-ozonated waters across a range of turbidities clearly indicate a strong benefit from ozone, with UVT values increasing by as much as 8 percentage points. True UVT values measured by integrated sphere also noted a significant benefit of ozonation, although this diminished with increasing turbidity in Coquitlam 2 water.

Impact of UVT measurement methods In light of the difference between direct and true UVT values, the results reported above need to be revisited. It was found that 4-log inactivation of MS2 and 3-log inactivation of Cryptosporidium could be achieved in 50 NTU water. The exposure times used were calculated incorporating the direct UV absorbance of the turbidity-adjusted water, a value that would be subject to the light scattering error. Substituting the true UVT for the direct UVT, as measured at the time of the collimated beam tests, the UV doses actually applied can be calculated, and are shown in Table 14.9. For the sake of completeness, this same assessment should be made of the UV dose applied to Cryptosporidium in elevated turbidity. The true applied dose would increase by the same proportion, but as Cryptosporidium is so sensitive to UV, the net effect of using direct UV in determining exposure time necessary for a target dose of 3.0 mJ/cm2 was to increase the actual applied dose to the 50 NTU sample up to 3.2 mJ/cm2.

Summary and Conclusions This project involved the seeding of surrogate or actual pathogenic organisms to low turbidity Coquitlam source water, determining their inactivation kinetics, and

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Table 14.9. UV doses applied to coliphage MS2 corrected for true UV absorbance.

Test water Turbidity (NTU) Baseline 5 10 15 20 50

Coquitlam 1 Targeted: 83 mJ/cm2

Coquitlam 2 Targeted: 92 mJ/cm2

True dose applied (mJ/cm2)

True dose applied (mJ/cm2)

83.0 83.0 83.0 83.0 83.6 85.3

92.0 92.0 92.5 93.3 94.0 98.5

The applied dose values obtained after making the correction for true absorbance (and UVT) are only notably different from the targeted dose for those tested at the highest turbidities.

then applying the disinfectant dose necessary for target inactivation to Coquitlam water that been adjusted with native particulate matter to a range of turbidity levels. In these latter tests, an effort was made to allow the test microorganisms to associate, by passive contact, with Coquitlam particulate matter, although no active flocculation steps were included. In addition to the evaluation of direct impacts to UV and ozone disinfection by turbidity, the ability of ozone to enhance the UV treatability of Coquitlam water was assessed as turbidity was artificially elevated. The results of this project have confirmed that GVRD has selected an effective multi-barrier approach for protecting the health of consumers of unfiltered drinking water. However, caution is recommended if any circumstances arise such that turbidity spikes due to some cause that enriches the organic rather than inorganic fraction of the suspended particles. An algal bloom or terrestrial upset, such as the immediate fallout of a forest fire, might have an effect not tested in this study. The protected nature of the watershed makes a nutrient influx fuelling an algal bloom an unlikely scenario, but a natural or human-induced forest fire is possible. Ozone was shown to be a very effective pretreatment of Coquitlam water for UV, regardless of turbidity. The UV transmittance of Coquitlam water, and thus its UV treatability, was uniformly improved by ozonation.

References Adams, M.H. (1959) Bacteriophages. Interscience Publishers, New York. Bolton, J.R. and Linden, K.G. (2003) Standardization of methods for fluence (UV dose) determination in bench-scale UV experiments. ASCE Journal of Environmental Engineering 129, 209–215. Clancy, J.L., Marshall, M.M., Hargy, T. and Korich, D.G. (2004) Susceptibility of five strains of Cryptosporidium parvum oocysts to UV light. Journal of the American Water Works Association 96, 84–93.

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T.M. Hargy et al. Dow, S.M., Barbeau, B., von Gunten, U., Chandrakanth, M., Amy, G. and Hernandez, M. (2006) The impact of selected water quality parameters on the inactivation of Bacillus subtilis spores by monochloramine and ozone. Water Research 40, 373–382. Health Canada (2005) Canadian Guidelines for Drinking Water Quality. Available at: http://www.hc-sc.gc.ca/ewh-semt/pubs/water-eau/doc_sup-appui/protozoa/ chap_12_e.html. Hoff, J.C., Rice, E.W. and Schaefer, F.W., III (1985) Comparison of animal infectivity and excystation as measures of Giardia muris cyst inactivation by chlorine. Applied and Environmental Microbiology, 50, 1115–1117. Meng, Q.S. and Gerba, C.P. (1996) Comparative inactivation of enteric adenovirus, poliovirus and coliphages by ultraviolet irradiation. Water Research 30, 2665–2668. Shin, G.-A., Linden, K.G., Arrowood, M.J. and Sobsey, M.D. (2001) Low-pressure UV inactivation and DNA repair potential of Cryptosporidium parvum oocysts. Applied and Environmental Microbiology 67, 3029–3032. USEPA (1992) Consensus Method for Determining Groundwaters under the Direct Influence of Surface Water using Microscopic Particulate Analysis (MPA). EPA 910992029. USEPA Environmental Services Division, Port Orchard, WA. Government Printing Office, Washington, DC. USEPA (2005) Method 1623: Cryptosporidium and Giardia in Water by Filtration/IMS/ FA. EPA 815-R-05-002. United States Environmental Protection Agency, Office of Water, Washington, DC. USEPA (2006) Ultraviolet Disinfection Guidance Manual for the Final Long Term. 2. Enhanced Surface Water Treatment Rule. Office of Water (4601) EPA 815-R-06-007, United States Environmental Protection Agency, Washington, DC. Wilson, B.R., Roessler, P.F., Van Dellen, E., Abbaszadegan, M. and Gerba, C.P. (1992) Coliphage MS-2 as a UV water disinfection efficacy test surrogate for bacterial and viral pathogens. In: Proceedings of the American Water Works Association Water Quality Technology Conference, Toronto, Canada. American Water Works Association, Denver, CO, USA.

15

Towards Methods for Detecting UV-induced Damage in Individual Cryptosporidium parvum and Cryptosporidium hominis Oocysts by Immunofluorescence Microscopy

H.V. SMITH1, B.H. AL-ADHAMI1, R.A.B. NICHOLS1, J.R. KUSEL2 AND J. O’GRADY3 1Scottish

Parasite Diagnostic Laboratory, Glasgow, UK; 2Institute of Biomedical and Life Sciences, Glasgow, UK; 3Strathclyde Institute of Pharmacy and Biomedical Sciences, University of Strathclyde, UK

Abstract Water is an important transmission route for cryptosporidiosis, with at least 165 waterborne outbreaks of cryptosporidiosis documented. Cryptosporidium can be controlled in water treatment by physical removal and UV disinfection, and a method that can determine whether individual oocysts in a routine sample exposed to UV irradiation have been disinfected is of benefit as it offers increased confidence to water operators. The major effects of UV radiation on cell membranes are alterations of proteins, particularly protein crosslinking. UV-B radiation progressively inhibits protein synthesis. Specific free radical scavengers protect cells against killing and inhibition of protein synthesis by UV-B. UV light also crosslinks the complementary strands of DNA and causes the formation of single strand breaks and pyrimidine dimers. The major lesions induced are cyclobutyl pyrimidine dimers (CPDs; also known as thymine dimers, TD). UV-induced DNA lesions in living cells and in some microorganisms can be repaired by the enzyme-dependent nucleotide excision repair (NER), also named dark repair, and the light-dependent reaction known as photoreactivation (PHR). Dark repair and PHR enable UV-inactivated microorganisms to recover and may reduce the efficiency of UV inactivation. Cryptosporidium parvum oocysts are inactivated at 3–40 mJ/cm2 using medium- and low-pressure UV light. Cryptosporidium parvum can undertake photoreactivation and dark repair at the genomic level and NER repair genes have been identified in C. parvum and C. hominis. However, UV inactivation of Cryptosporidium oocysts is irreversible, despite the presence of the UV repair genes. © CAB International 2009. Giardia and Cryptosporidium: From Molecules to Disease (eds G. Ortega-Pierres et al.)

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H.V. Smith et al. We investigated the following hypotheses for developing a method to demonstrate UV inactivation of Cryptosporidium oocysts: (i) UV disinfection induces the production of reactive oxygen species (ROS); (ii) UV disinfection induces apoptosis; and (iii) UV disinfection causes damage to DNA, which is detectable using fluorogenic DNA reporters. We developed assays for determining UV inactivation of Cryptosporidium oocysts which would remain compatible with, and integrated as much as possible with, existing UK and USA detection methodologies. Our development of suitable methods which can detect UV damage in individual organisms reliably and reproducibly was driven by a search for fluorogenic reporters which could enter and stain UV-killed and damaged Cryptosporidium oocysts. Antioxidants reduce ROS production and the antioxidant glutathione (GSH) plays a significant role in inhibiting the generation of mutagens by ionizing radiation. The presence of GSH, as a putative reporter of UV damage caused by the production of ROS, was investigated in intact, untreated C. parvum oocysts and sporozoites within intact UV-irradiated oocysts (40 mJ/cm2) using the fluorogenic vital dye monochlorobimane (MCB) to detect both GSH levels and activity in oocysts. MCB fluorescence localized GSH in purified, intact, recently excreted and aged C. parvum oocysts, at several nuclear and cytoplasmic sporozoite foci (n = 2–6). We did not demonstrate the function of GSH as an endogenous free radical scavenger in UV-irradiated oocysts, and other free radical scavengers are more active than GSH in UV-treated C. parvum oocysts. MCB is unlikely to be useful as a surrogate for detecting UV damage in UV-treated Cryptosporidium oocysts. The DNA intercalating dye YO-PRO1 (YP) has been used to determine apoptosis and was used to investigate the role of UV irradiation in inducing programmed cell death/ apoptosis. YP detected DNA damage in UV-treated (40 mJ/cm2) C. parvum oocysts. YP was incorporated into sporozoite DNA of intact, irradiated oocysts (possibly apoptotic) which exhibited no apparent oocyst wall damage. However, control oocysts did not exclude YP entirely. YP is unlikely to provide a reliable estimate of the possible apoptotic changes that can occur in irradiated oocysts. An antibody raised against TDs (α-TD) was used to identify changes induced by UV light in C. parvum sporozoites and oocysts, and its nuclear location was validated by co-localization with DAPI. A freeze-thawing (five cycles) procedure improved α-TD antibody labelling within irradiated C. parvum oocysts. No α-TD localization was seen in non-irradiated oocysts. Both C. parvum and C. hominis oocysts exposed to different doses of UV light (range 10–40 mJ/cm2) demonstrated TD lesions following irradiation. We conclude that an immunofluorescence assay using α-TD antibodies which, for C. parvum, has been validated against a neonatal mouse infectivity assay, is suitable for detecting thymine dimers in air-dried oocysts and air-dried sporozoites of C. parvum and C. hominis oocysts, and that the α-dsDNA antibody is a good candidate for a positive control for the assay.

Introduction Cryptosporidium oocysts are frequent contaminants of water with contributions from infected human and non-human hosts, livestock and agricultural practices, and infected feral and transport hosts (Smith and Rose, 1990, 1998; Smith et al., 1995). Recent genetic analyses have raised doubt about the validity of the current classification of the genus Cryptosporidium and reveal that more than one species of Cryptosporidium can infect susceptible, immunocompetent

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human hosts (Table 15.1). There are 16 ‘valid’ Cryptosporidium species and a further 40+ genotypes, which differ significantly in their molecular signatures but, as yet, have not been ascribed species status (Smith et al., 2007). Seven described Cryptosporidium species (C. hominis, C. parvum, C. meleagridis, C. felis, C. canis, C. suis and C. muris) (Table 15.1) and two undescribed species of Cryptosporidium (cervine and monkey) infect immunocompetent and immunocompromised humans (Xiao et al., 2004; Cacciò et al., 2005), but C. hominis and C. parvum are the most commonly detected (Cacciò et al., 2005). Water is an important transmission route, with at least 165 waterborne outbreaks of cryptosporidiosis documented (Girdwood and Smith, 1999; Fayer et al., 2000; Slifko et al., 2000; Karanis et al., 2007). Cryptosporidium can be controlled in water treatment plants by physical removal and disinfection processes. Physical characteristics, such as size, settling velocities and surface charge affect the behaviour of oocysts in physical treatment processes (Smith et al., 1995). Essentially, oocysts behave as inert, discrete particles in water. Current data indicate that Cryptosporidium is controllable in filtration processes and the performance of individual treatment plants in removing oocysts may be expected to parallel their effectiveness for turbidity control. The major factors that influence a plant’s performance for turbidity control should be expected to affect the efficiency of removal of Cryptosporidium similarly. Cryptosporidium oocysts are insensitive to commonly used disinfectants (chlorine, chlorine dioxide, ozone, etc.), but UV disinfection is effective. Clearly, a method that can determine whether all oocysts in a routine sample exposed to UV irradiation have been disinfected is of benefit, as it offers increased confidence to water operators. Here, we focus on UV disinfection of Cryptosporidium oocysts. Oocysts occur at low densities in water (Smith and Rose, 1990, 1998; Smith et al., 1995) and methods which can detect and determine the UV sensitivity of small numbers of organisms reliably and reproducibly from water concentrates are required. Little can be inferred about the likely impact of oocysts detected in water concentrates on public health without knowing whether they are viable or not. The conventional techniques of animal infectivity and excystation in vitro are not applicable Table 15.1.

Some differences among Cryptosporidium species infecting humans.

Species C. hominis C. parvum C. suis C. felis C. canis C. meleagridis C. muris C. andersoni

Oocyst dimensions (µm)

Site of infection

Major host

4.5 × 5.5 4.5 × 5.5

Small intestine Small intestine

5.05 × 4.41 4.5 × 5.0 4.95 × 4.71 4.5–4.0 × 4.6–5.2 5.5 × 7.4 5.6 × 7.4 (5.0–6.5 × 8.1–6.0)

Small intestine Small intestine Small intestine Intestine Stomach Stomach

Humans Neonatal mammalian livestock, humans Pigs Cats Dogs Turkeys Rodents Cattle

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to the small numbers of organisms found in water concentrates, and much effort has been expended on the development of surrogate techniques that can address, accurately, the viability of individual oocysts. Fluorogenic vital dye methods are based on observing whether specific fluorogenic vital dyes are included into or excluded from Cryptosporidium oocysts as a measure of their viability.

The Usefulness of Existing Fluorogenic Vital Reporters for Determining Disinfection Capability is Compromised A variety of fluorescent dyes have been used to determine Cryptosporidium oocyst viability. In a joint UK/USA initiative to test in vitro (maximized in vitro excystation, fluorogenic vital dyes assay (4′,6-diamidino-2-phenylindole (DAPI) and propidium iodide (PI); DAPI-PI), SYTO9 and SYTO59) and in vivo (neonatal CD-1 mouse infectivity) surrogates (Clancy et al., 2000), high levels of variability were detected with both sets of surrogates for untreated, aged and disinfectant-treated controls. Variability in in vitro surrogates was probably associated with subsampling errors (enumeration of only 100 oocysts from a large oocyst stock), while variability in the in vivo surrogate was attributed to delivery of a standard dose by gavage and the use of outbred animals. In vitro assays overestimated oocyst viability compared with neonatal mouse infectivity, particularly following pulsed UV disinfection. Overall, maximized in vitro excystation and SYTO9 produced closer association with mouse infectivity than DAPI-PI. SYTO59 produced the greatest disparity between in vitro and in vivo assays (Clancy et al., 2000). In the UV treatment studies, SYTO9 and SYTO59 consistently demonstrated higher oocyst viabilities than maximized in vitro excystation and DAPI-PI, even though mouse infectivity failed to demonstrate infectious oocysts. The majority of published data indicate that while fluorogenic vital dyes can be useful predictors of oocyst viability, they also underestimate the degree of oocyst inactivation following chemical disinfection (Bukhari et al., 1999; Clancy et al., 2000; O’Grady and Smith, 2002). Thus, studies evaluating the effectiveness of chemical disinfectants using in vitro viability procedures may have reported higher than necessary disinfection requirements due to the overestimation of viability compared with in vivo procedures.

UV Light Disinfection UV light is classified according to wavelength: UV-A, 315–400 nm; UV-B, 280–315 nm; and UV-C, 100–280 nm; and has been used for drinking water disinfection since the beginning of the 20th century. UV light disinfection does not generate significant disinfection by-products. It does not cause a significant increase in assimilable organic carbon, nor does it convert nitrates to nitrites, nor bromide to bromines or bromates. UV disinfection is relatively insensitive to

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temperature and pH differences. Disadvantages include its unsuitability for water with high levels of suspended solids, turbidity, colour, or soluble organic matter, as they can react with UV radiation, reducing disinfection performance. Turbidity reduces the water penetration of UV radiation and there is no disinfection residual.

Mode of action of UV light disinfection The modes of action of UV light on uni- and multicellular organisms involve oxidative stress resulting from the attack by free radicals of several cellular targets (proteins, DNA and lipids). They are primarily twofold, as follows: 1. The major effects of UV radiation on cell membranes are alterations of proteins, particularly protein crosslinking (detected as high molecular weight protein >200 kDa) (Kochevar, 1990). UV light can also crosslink exposed collagen, making it more resistant to enzymic digestion and less elastic (Lee et al., 2001). UV-B radiation progressively inhibits protein synthesis and kills Staphylococcus aureus. The OH- and 1O2-free radical scavengers protect cells against killing and inhibition of protein synthesis by UV-B, suggesting that such radicals mediate the effects of UV-B on this organism. A similar protective effect using a ferric ion chelator suggests an important role for metallic ions in UV-B lethality (El-Adhami et al., 1994). Additional effects of pulsed UV light damage include increased concentration of eluted proteins, increased cell membrane damage, and structural changes in yeast cells (Takeshita et al., 2003). 2. UV light crosslinks the complementary strands of DNA. It also causes the formation of single strand breaks and pyrimidine dimers (Takeshita et al., 2003). DNA conformation and/or flexibility governs the phenomenon of crosslinking. (GA).(TC) suppresses the crosslink formation in DNA more than any dinucleotide composed of only G and C. (CTAG).(CTAG) promotes crosslinking much more than any other tetranucleotide, including (TATA).(TATA), whereas the closely related (CATG).(CATG) belongs among the tetranucleotides that most suppress the UV-light-induced crosslinks between the complementary strands of DNA (Nejedly et al., 2001a). Nejedly et al. (2001b) also identified that the (ATTTTATA).(TATAAAAT) octamer is a candidate for the hotspot of UV-light-induced crosslinking between the complementary strands of DNA. UV light damages DNA, and the major lesions induced are cyclobutyl pyrimidine dimers (CPDs; also known as thymine dimers, TD) (Mitchell, 1988). The UV-induced DNA base modifications lead to the production of reactive oxygen species (ROS), which can damage cellular elements. The antioxidant glutathione (GSH) plays a significant role in inhibiting the generation of ROS (Fischer-Nielsen et al., 1994). UV-induced DNA lesions in living cells (Roza et al., 1991) and in some microorganisms (Oguma et al., 2001) can be repaired by one or more mechanisms. Such mechanisms include the enzyme-dependent nucleotide excision repair (NER), also named dark repair, and the light-dependent reaction

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known as photoreactivation (PHR). Dark repair and PHR enable UV-inactivated microorganisms to recover and may reduce the efficiency of UV inactivation. Some microorganisms found in water exhibit light and/or dark repair.

UV light and disinfection of Cryptosporidium oocysts The potential of UV irradiation for disinfecting drinking water has been recognized for many years. UV irradiation is effective for killing microorganisms, including bacteria, fungi and oocysts. Various studies have shown that parasitic protozoa are more sensitive to UV irradiation than viruses, but less sensitive than most bacteria (reviewed in Hijnen et al., 2006). Cryptosporidium parvum oocysts can be inactivated using pulsed UV light (Slifko et al., 1999) and low- (Oguma et al., 2001; Shin et al., 2001) and medium-pressure UV (Craik et al., 2001; Rochelle et al., 2004). Pulsed light consists of intense flashes of broad-spectrum white light containing wavelengths from 200 nm (UV) to 1000 nm in the near infrared region (Takeshita et al., 2003). Low-pressure mercury lamps emit peak output at 254 nm, whereas medium-pressure mercury lamps emit radiation at several peaks between 248 and 295 nm. Most studies on C. parvum oocyst inactivation have used low- and medium-pressure lamps (Rochelle et al., 2005). No differences in UV inactivation of C. parvum are apparent using either low- or medium-pressure UV lamps (Craik et al., 2001; Rochelle et al., 2004). Table 15.2 identifies some studies that indicate that UV light is an effective disinfectant for waterborne Cryptosporidium oocysts, particularly when mediumand low-UV-pressure lamps are used. The majority of data on the usefulness of UV disinfection have been accrued using sources which deliver continuous UV irradiation; however, the possibility that pulsed UV light may be more effective for water disinfection has also been put forward (Takeshita et al., 2003).

Assays for Determining UV Inactivation of Cryptosporidium Oocysts A variety of assay methods using a variety of C. parvum oocyst isolates have been used. In one of the earliest published works, C. parvum oocyst inactivation by UV radiation was assessed using maximized in vitro excystation and the fluorogenic vital dyes assay. Oocysts were inactivated (>99%, 2-log10 reduction) using a high dose of UV irradiation (up to 8748 mJ/cm2) (Campbell et al., 1995). Morita et al. (2002) showed that C. parvum oocysts exhibited high resistance to UV irradiation, requiring a dose of 230 mJ/cm2 for a 99% reduction in excystation. Animal infectivity and in vitro cell culture studies indicated that C. parvum oocysts were much more sensitive to UV irradiation than previously identified: the UV dose required for a 99% reduction in mouse infectivity was 1.0 mJ/cm2, whereas a dose of approximately 200 mJ/cm2 was required to achieve 99% reduction by excystation (Morita et al., 2002). Rochelle et al. (2004) showed that 99.9% inactivation (0.001% relative mouse infectivity) was achieved with a UV dose of 7.5 mJ/cm2. Using the CD-1 neonatal mouse infectivity assay, 99.9%

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Table 15.2. UV disinfection of Cryptosporidium oocysts determined by oocyst inactivation. UV pressure

UV dose

Time/mode

Log reduction

Reference

Low

120 mJ/cm2

4 gpm/water device

Drescher et al. (2001)

Not stated

15,000 mW/sec = 15,000 mJ/ cm2

150 min/ bench scale

Medium and low

0.8–119 mJ/cm2

Bench scale

Medium

41 mJ/cm2

Bench scale

Low

20–10 mJ/cm2

Bench scale

5.4 with neonatal mouse infectivity >2 with neonatal mouse infectivity Max. 3.4–4.9 with neonatal mouse infectivity >4 with neonatal mouse infectivity >3 with in vitro infectivity

LorenzoLorenzo et al. (1993) Craik et al. (2001)

Bukhari et al. (1999)

Bukhari and LeChevallier (2003)

inactivation at a UV dosage of 19 mJ/cm2 (Bukhari et al., 1999) and 99.0–99.9% inactivation at 10–25 mJ/cm2 (Craik et al., 2001) could be achieved. Shin et al. (2001) utilized a cell culture infectivity assay combined with epifluorescence detection to determine UV inactivation of C. parvum. They reported 99.9% inactivation at a dosage of 3 mJ/cm2 using a low-pressure UV lamp. Both cell culture and animal infectivity assays gave comparable results with a 99.9% inactivation at a dosage of 3 mJ/cm2. Rochelle et al. (2004) assessed the inactivation of different isolates of C. parvum exposed to low- or medium-pressure UV lamps, using cell culture (monolayers of HCT-8 cells) and a reverse transcriptase polymerase chain reaction (RT-PCR) assay. An average dose of 7.6 mJ/cm2 resulted in 99.9% inactivation of oocysts in five different isolates. Using the same culture-based methods, Johnson et al. (2005) demonstrated that C. hominis display similar levels of infectivity in cell culture and have a similar sensitivity to UV light as C. parvum. Cryptosporidium parvum can undertake photoreactivation and dark repair at the genomic level (Oguma et al., 2001; Morita et al., 2002). In addition, NER repair genes have been identified in C. parvum and C. hominis. However, UV inactivation of Cryptosporidium oocysts is irreversible, despite the presence of the UV repair genes (Rochelle et al., 2004).

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Application to Waterborne Oocysts In order to remain compatible with, and to integrate as much as possible with, existing UK and USA detection methodologies, our development of suitable methods that can detect UV damage in individual organisms reliably and reproducibly was driven by a search for fluorogenic reporters which will enter and stain UV killed and damaged Cryptosporidium oocysts. This was guided by the following hypotheses: 1. UV disinfection induces the production of reactive oxygen species (ROS). 2. UV disinfection induces apoptosis. 3. UV disinfection causes damage to DNA, which is detectable using fluorogenic DNA reporters.

Hypothesis 1: UV induces the production of reactive oxygen species (ROS) Ultraviolet light A (UV-A) radiation induces the production of ROS in cells, which can damage cellular elements. Oxidative modifications to DNA nucleotides (e.g. 8-hydroxyguanine (8-oxoG)) are mutagenic (Cheng et al., 1992). Antioxidants reduce ROS production and GSH plays a significant role in inhibiting the generation of 8-oxoG by ionizing radiation (Fischer-Nielsen et al., 1994). GSH reacts with various ROS and is a cofactor for the H202-removing enzyme, glutathione peroxidase. In cultured mammalian cells, GSH depletion and thermal stress increase endogenous oxidative damage, but the addition of thiols to the medium does not reduce the level of oxidative damage caused by GSH depletion and thermal stress (Will et al., 1999). GSH is a low-molecular-weight tripeptide (gamma-glutamylcysteinylglycine) which is synthesized intracellularly. It plays a critical role in the detoxification of several drugs and xenobiotics (Meister and Anderson, 1983) and in cellular defence against agents that cause oxidative stress (Anderson, 1998). A very well-established technique to measure GSH is to add the cell permeant, fluorogenic vital dye monochlorobimane (MCB) to detect both GSH levels and activity in cells. MCB does not fluoresce, but on reacting with GSH yields GSH-bimane adducts which fluoresce. The enzyme glutathioneS-transferase (GST) exclusively mediates the intracellular conjugation of GSH and MCB. The rate of conjugation between GSH and MCB (which produces the fluorescence signal) is dependent on the abundance of GST (Haugland, 2005). As a putative reporter of UV damage caused by the production of ROS, we investigated the presence of the antioxidant GSH in C. parvum sporozoites in intact, untreated oocysts and in sporozoites within intact UV-irradiated oocysts (40 mJ/cm2). Purified C. parvum oocysts were purchased from Bunch Grass Farm (Idaho, USA; Iowa isolate BGF06-1, 10 days old) and stored between 4°C and 8°C until used. The percentage viability and excystation rate of this isolate using our optimized, fluorogenic vital dyes (DAPI-PI; Campbell et al., 1992) and our maximized in vitro excystation protocols (Robertson et al., 1993) were 95.2 ± 2.3 and 96.0 ± 1.0, respectively.

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UV irradiation was delivered using a high-intensity, low-pressure UV lamp with an output at 254 nm. The procedure was adapted from the method described by Rochelle et al. (2004). Short-wave UV irradiation from a UVGL-58 Mineralight lamp was utilized. The intensity of the UV light, measured using a digital UVX radiometer, was (on average) 350 µW/cm2 at 254 nm. A rig was set up, and 10 cm below the lamp a position was marked where the UV intensity was maximal (350 µW/cm2). The UV dose was then determined from: UV dose (mJ/cm2) = Irradiance (mW/cm2) × Exposure time (s) In all experiments, 1 × 106 oocysts were suspended in 5 ml of Hanks’ balanced salt solution (HBSS). Samples were placed in Petri dishes (diameter 36 mm) which were constantly mixed using a magnetic stirrer during exposure to UV light. To achieve different UV dosages (mJ/cm2), oocysts were exposed to UV light for varying times at a constant distance (10 cm) from the constant intensity UV source. For each experiment, control oocysts were kept under the same conditions without irradiation. Statistical analysis was performed using analysis of variance (ANOVA) with P < 0.05 as the criterion for significance using MINITAB version 11 software. MCB fluorescence localized GSH in intact oocysts and the number of distinctly labelled foci in intact oocysts varied between 2 and 6 both in recently excreted (10 days old) and aged (6 months old) oocysts (Al-Adhami et al., 2006). MCB-labelled excysted sporozoites retained the fluorogenic dye at several intrasporozoite foci. GSH distribution in sporozoites was granular in the apical and posterior (nuclear) regions (Al-Adhami et al., 2006). We used buthionine sulphoximine (BSO), a potent and specific inhibitor of GSH (Meister and Anderson, 1983), to determine whether GSH is synthesized in BSO-treated oocysts, by labelling treated oocysts with MCB. When oocysts were depleted of GSH using BSO for 24 h at room temperature (RT), a significant decrease in fluorescence (~50%) was observed, indicating that MCB binds sporozoite GSH. We enumerated oocysts exposed to 10, 20 or 40 mJ/cm2 doses of UV light for MCB inclusion or exclusion, and quantified their fluorescence (using ANALYSIS software for scientific imaging and calibrated image measurements; Olympus, UK). The percentage inhibition of irradiated versus control oocysts was not significantly different (irradiated group mean ± SD 19.8 ± 10.5, 17.2 ± 12.8, 20.6 ± 8.0 at 10, 20 or 40 mJ/cm2, respectively; control group mean ± SD 15.5 ± 9.6, P > 0.05). No significant differences in fluorescence intensity or distribution of MCB occurred in irradiated or control oocysts (irradiated group mean ± SD mJ/cm2 = 152.3 ± 15.9, 20 mJ/cm2 = 149.6 ± 12.2, 40 mJ/ cm2 = 160.2 ± 18.5; control group mean ± SD 165.1 ± 20.3, P > 0.05). Oocysts subjected to GSH depletion showed significant decrease in GSH-bimane adducts (mean ± SD 62.3 ± 13.4) when compared to the control group (mean ± SD 165.1 ± 20.3, P = 0) as demonstrated by fluorescence quantitization.

Determining oocyst viability using MCB and propidium iodide staining The procedure developed was similar to that used in the fluorogenic vital dyes (DAPI-PI) assay of Campbell et al. (1992). The proportions of MCB+

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PI−, MCB− PI+, MCB− PI− and empty oocysts were quantified by enumerating 100 oocysts in triplicate samples. Oocysts were considered viable if they did not include PI but were stained with MCB (MCB+ PI−). Also, intact oocysts which did not include MCB or PI (MCB− PI−) but contained sporozoites under differential interference contrast (DIC) microscopy were considered viable. MCB incorporation correlated well with DAPI+ PI− oocyst staining. The results of DAPI-PI and MCB-PI staining were not significantly different when labelled oocysts were enumerated (mean ± SD 84.7 ± 4.2 versus 73.3 ± 8.7, P = 0.05, respectively). Results using the maximized in vitro excystation assay were not significantly different from DAPI-PI staining (mean ± SD 89.7 ± 2.1 versus 84.7 ± 4.2, P = 0.1) but were significantly different from MCB-PI staining (mean ± SD 89.7 ± 2.1 versus 73.3 ± 8.7, P = 0.01). These data, based on the MCB labelling of intact C. parvum oocysts, identify the presence of glutathione both in nuclear and cytoplasmic foci of sporozoites, which can be specifically depleted by BSO. Its function as an endogenous free radical scavenger in UV-irradiated oocysts was not demonstrated. Thus it is likely that other free radical scavengers are more active than GSH in UV-treated C. parvum (e.g. cysteine, ascorbic acid). MCB is unlikely to be useful as a surrogate for detecting UV damage in UV-treated Cryptosporidium oocysts.

Hypothesis 2: UV disinfection induces apoptosis The role of UV irradiation in inducing programmed cell death/apoptosis was investigated using DNA intercalating dyes. Currently, there are no reports available regarding UV-irradiation-induced apoptotic changes in Cryptosporidium oocysts. YO-PRO1 (YP) binds strongly to nucleic acids and has been used to detect apoptosis in mammalian cells without interfering with cell viability (Idziorek et al., 1995). Apoptosis is a highly regulated physiological process that is linked to pathological events such as oxidative stress in which the reactive oxygen species (ROS) play a key role in the initiation of the process (Plantin-Carrenard et al., 2003). The early stages of apoptosis are characterized by chromatin condensation, nuclear fragmentation and mitochondrial clustering, but with no loss of membrane integrity (Cohen, 1993). The plasma membrane becomes slightly permeable during apoptosis, allowing the uptake of YP but not PI, the impermeant dead-cell stain (Gilbert and Knox, 1997). Apoptosis is accompanied by the activation of an endonuclease enzyme that cleaves DNA initially into large fragments (50–300 kb) which yield internucleosomic fragments of 180–200 base pair multimers (Cohen et al., 1994). YP nucleic acid stain forms the basis of an important assay for apoptotic cells and is compatible with both epifluorescence microscopy and flow cytometry. Selective uptake of YP by apoptotic cells of a dexamethasone-treated population of thymocytes, an irradiated peripheral blood mononuclear cell population, and a growth-factor-depleted tumour B cell line was confirmed by cell sorting (Idziorek et al., 1995). Apoptotic cells take up YP, while viable cells exclude it (Haugland, 2005).

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We focused our investigations on the usefulness of YP to detect DNA damage in UV-treated (40 mJ/cm2) C. parvum oocysts. Three nucleic acid stains were used to identify viable (DAPI, blue nuclei; in the absence of PI, red nuclei) oocysts, apoptotic (YP, green nuclei) oocysts and dead (PI, red nuclei) oocysts. UV disinfection procedures were as previously described. Epifluorescence microscopy, flow cytometry and Nomarski differential interference contrast (DIC) microscopy were used to assess outcomes. Oocysts exposed to UV disinfection, those subjected to apoptosis-inducing drug treatment changes, and controls were analysed using this three-colour assay. DAPI (indicator of viability in the absence of PI staining) was taken up by ~95% of the untreated population, while YP stained 35% of the same population. PI was excluded from all DAPI+ YP+ oocysts. PI stained the remaining 5% of the population. This result was consistent with that obtained with the DAPI-PI assay. YP was used in combination with PI to assess the population structure of irradiated, drug-treated and untreated oocysts by epifluorescence microscopy and fluorescence activated cell sorting (FACS). YP was incorporated into sporozoite DNA of intact, irradiated oocysts (possibly apoptotic) which exhibited no oocyst wall damage as determined by DIC microscopy. Apoptosis was induced in oocysts treated with dexamethasone (1 mM) or etoposide (10 µM) prior to labelling with fluorescent probes. Drug treatment induced increased YP signals compared with controls. However, control oocysts did not exclude YP entirely, showing low-level staining. DIC microscopy revealed alterations in the morphology of drug-treated oocysts, compared with irradiated and control (untreated) oocysts. When oocysts were identified as apoptotic (YP+ DAPI+ PI−), no significant differences were detected between irradiated and drug-treated oocysts, compared with controls (Table 15.3). Unlike our epifluorescence microscopy data, FACS revealed that 71.9% of irradiated and 33.7% of drug-treated populations were YP-positive compared with 4.2% of the control population. Despite the increase in YP-positive oocysts by FACS, it is unlikely that YP can provide a reliable estimate of the possible apoptotic changes that can occur in irradiated oocysts or sporozoites, for two main reasons: (i) the staining pattern resulting from the simultaneous use of these Table 15.3. Epifluorescence microscopy and flow cytometric analyses of recently excreted C. parvum oocysts subjected to different treatments, then stained with YP, DAPI and PI for epifluorescence microscopy or YP and PI for flow cytometry. % of labelled oocysts Assay Epifluorescence microscopy (YP+DAPI+PI−) (YP−DAPI+PI−) (YP−DAPI−PI+) Flow cytometry

Irradiated

Etoposide

Dexamethasone

Control

42.2 54.0 3.8 71.9

35.6 54.1 10.3 ND

ND ND ND 33.7

39.3 57.3 3.4 4.2

ND: Not determined. Note: YP stains apoptotic oocysts, PI stains dead oocysts.

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three dyes makes it impossible to distinguish normal from apoptotic oocysts by standard sets of filters used in epifluorescence microscopy; and (ii) differences in the intensity and the rapid quenching of YP make it very difficult to count and photograph YP-labelled oocysts.

Hypothesis 3: UV disinfection causes damage to DNA, which is detectable using fluorogenic DNA reporters The major DNA lesion caused by UV, the cyclobutyl pyrimidine dimer (TD), is responsible for UV-induced cytotoxicity and mutagenicity in living cells and microorganisms (Mitchell, 1988), and the formation of such lesions in genomic DNA inhibits normal replication and transcription of DNA and results in the inactivation of cells. Two known antibody reporters of nuclear UV damage were investigated. Two different antibodies raised against nucleic acids (α-double sDNA (α-dsDNA) and α-TD) were used to identify changes induced by UV light in C. parvum sporozoites and oocysts. Sporozoite nuclei were also stained with DAPI to validate the co-localization of α-dsDNA and α-TD (Al-Adhami et al., 2007). An improved α-TD antibody labelling procedure within irradiated C. parvum oocysts was established following freeze-thawing (five cycles), based on that of Nichols and Smith (2004). No α-TD localization was seen in non-irradiated oocysts. α-dsDNA antibody bound to the nuclei of both irradiated and non-irradiated sporozoites. Both C. parvum and C. hominis oocysts exposed to different doses of UV light (range 4–40 mJ/cm2) were tested using a standardized set of parameters: oocysts dried onto slides prior to freeze-thawing (five cycles), then fixed in methanol and labelled with either α-TD-Ab, a commercially available, fluorescein-labelled, monoclonal antibody reactive with surface exposed epitopes on Cryptosporidium oocysts (FITC-C-mAb) and DAPI stain or α-TD-Ab and DAPI. Our data indicate that UV irradiation at doses ranging from 10 to 40 mJ/cm2 can be detected using α-TD-Ab and DAPI (Al-Adhami et al., 2007). While the combination of α-TD-Ab, FITC-C-mAb and DAPI produced positive outcomes only with high levels of UV irradiation (40 mJ/cm2), if we replaced FITC-C-mAb with Texas Red (TR)-C-mAb and used the combination of α-TD-Ab, TR-C-mAb and DAPI, DNA damage to C. parvum and C. hominis sporozoites within intact oocysts could be detected at a lower limit of 10 mJ/cm2, but not at 4 mJ/cm2. Currently, the combination of α-TD-Ab, TR-C-mAb and DAPI can be used to detect damage in nuclei of oocysts of C. parvum and C. hominis exposed to UV light (range 10–40 mJ/cm2). Validation by comparison with infectivity in neonatal CD-1 mice All procedures and manipulations performed were as described by Korich et al. (2000), and the experimental design is presented in Table 15.4. Mice infected with untreated oocysts (standard curve, Table 15.4) showed varying levels of infection. In the standard curve group, the percentage infection ranged from 5.2% in mice infected with 30 oocysts to 80% in mice infected with 300 oocysts.

Towards Methods for Detecting UV-induced Damage Table 15.4.

191

Experimental design of animal infectivity experiment.

UV dose (mJ/cm2) 10 10 20 20 None (control) None (control) None (control)

No. of animals 20 20 20 20 20 20 20

Dose (No. of oocysts/10 µl HBSS) 300/10 µl (low) 3000/10 µl (high) 300/10 µl (low) 3000/10 µl (high) 30/10 µl (low) 150/10 µl (medium) 300/10 µl (high)

UV-irradiated oocysts of C. parvum at 10 and 20 mJ/cm2 (Table 15.4) failed to cause infection in neonatal CD-1 mice, despite the fact that large doses of oocysts (up to 3000 per os, which is greater than 44 times the ID50 for this strain of mouse; Korich et al., 2000) were given to the animals. We conclude that an immunofluorescence assay using α-TD antibodies, which, for C. parvum, has been validated against a neonatal mouse infectivity assay, is suitable for detecting thymine dimers in air-dried oocysts and air-dried sporozoites of C. parvum and C. hominis oocysts and that the α-dsDNA antibody is a good candidate for a positive control for the assay.

Genotyping Cryptosporidium Oocysts by PCR Following UV Disinfection The species of waterborne Cryptosporidium oocysts recovered from routine Cryptosporidium monitoring of water sources can be determined by molecular methods (see Smith et al., Chapter 17, this volume). One pertinent issue that arises is whether oocysts which have been disinfected by UV light can be amplified by PCR, as UV disinfection damages DNA and the apoptopic cascade activates an endonuclease enzyme that cleaves DNA initially into large fragments (50–300 kb), which yield internucleosomic fragments of 180–200 base pair multimers (Cohen et al., 1994). Statements to the effect that UV light at up to 40 mJ/ cm2 damages DNA to such an extent that the molecular tools used to determine the species of small numbers of oocysts (nested 18S rRNA loci) cannot be used, have been made on numerous occasions. UV irradiation damage and repair is detectable by PCR in mammalian cultured cells and assay sensitivity is dependent on the size of the DNA fragment amplified. Long PCRs (6–24 kb) are required to ensure sensitive and unequivocal detection of DNA-induced lesions. UV irradiation induces DNA helix distortion, caused by the formation of pyrimidine dimmers, and DNA synthesis is impaired both in vivo and in vitro. Wang et al. (2003) demonstrated damage and repair of the p53 gene in human cells by a multiplex long quantitative PCR, designed to co-amply a 7 kb fragment of the gene and a 500 bp fragment control to increase the reliability of the assay. The lesion frequency detected in this gene

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was 0.63 lesions/10 kb/10 J/m2. In most applications, the UV dosage required to induce detectable lesions by PCR inhibition is much higher than the dosage used for Cryptosporidium inactivation; furthermore, the PCR fragment size is much larger than is used in most PCR applications. However, less PCR product was observed after amplifying the 1.7 kb 16S rDNA gene from cyanobacteria after UV-B radiation-induced damage (Kumar et al., 2004). Genomic DNA, extracted from bacteria exposed to 2.5 W/m2 of irradiation (up to 90,000 mw/s/ cm2 = 90,000 mJ/cm2), was tested by random amplification of polymorphic DNA (RAPD) analysis and direct PCR, and a marked decrease in amplification products, which was directly proportional to exposure time, occurred with both assays. Most PCR applications on the Cryptosporidium 18S rRNA gene used for genotyping oocysts from water do not exceed ~1000-bp fragment length for the first PCR amplification and vary from ~400 to 800 bp for the secondary amplification of nested assays. By extrapolation, it is unlikely that UV irradiation of water will have an adverse effect on the PCRs used for detecting oocysts; however, experimental confirmation of this hypothesis must be sought.

Conclusions Water is an important transmission route for cryptosporidiosis, with at least 165 waterborne outbreaks documented. Cryptosporidium can be controlled in water treatment by physical removal, yet Cryptosporidium oocysts are insensitive to commonly used disinfectants, with the exception of UV. Clearly, a method that can determine whether the small numbers of oocysts found in routine samples exposed to UV irradiation have been disinfected is of benefit, as it offers increased confidence of the success of UV disinfection to water operators. Neonatal animal infectivity, in vitro infectivity, excystation in vitro and current fluorogenic vital dye methods cannot be used to address, accurately, the viability of individual UV disinfected oocysts. UV light is classified according to wavelength: UV-A, 315–400 nm; UV-B, 280–315 nm; and UV-C, 100–280 nm; has been used for drinking water disinfection since the beginning of the 20th century, and is relatively insensitive to temperature and pH differences. Turbidity reduces its water penetration radiation and there is no disinfection residual. The modes of action of UV light on uni- and multicellular organisms involve oxidative stress resulting from the attack by free radicals of several cellular targets. The major effects of UV radiation on cell membranes are alterations of proteins, particularly protein crosslinking. UV-B radiation progressively inhibits protein synthesis. Specific free radical scavengers protect cells against killing and inhibition of protein synthesis by UV-B. Additional effects of pulsed UV light damage include increased concentration of eluted proteins, increased cell membrane damage and structural changes in yeast cells. UV light also crosslinks the complementary strands of DNA and causes the formation of single strand breaks and pyrimidine dimers. The major lesions induced are cyclobutyl pyrimidine dimers (CPDs; also known as thymine dimers, TD).

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UV-induced DNA lesions in living cells and in some microorganisms can be repaired by enzyme-dependent nucleotide excision repair (NER), also named dark repair, and the light-dependent reaction known as photoreactivation (PHR). Dark repair and PHR enable UV-inactivated microorganisms to recover and may reduce the efficiency of UV inactivation. C. parvum oocysts are inactivated at 3–40 mJ/cm2 using medium- and low-pressure UV light. C. parvum can undertake photoreactivation and dark repair at the genomic level, and NER repair genes have been identified in C. parvum and C. hominis. However, UV-inactivation of Cryptosporidium oocysts is irreversible, despite the presence of the UV repair genes. We developed assays for determining UV inactivation of Cryptosporidium oocysts which would remain compatible with, and integrated as much as possible with, existing UK and USA detection methodologies. Our development of suitable methods which can detect UV damage in individual organisms reliably and reproducibly was driven by a search for fluorogenic reporters which could enter and stain UV-killed and damaged Cryptosporidium oocysts, and was guided by the following hypotheses: 1. UV disinfection induces the production of reactive oxygen species (ROS). Antioxidants reduce ROS production and the antioxidant glutathione (GSH) plays a significant role in inhibiting the generation of mutagens by ionizing radiation. We investigated the presence of GSH, as a putative reporter of UV damage caused by the production of ROS, in intact, untreated C. parvum oocysts and sporozoites within intact UV-irradiated oocysts (40 mJ/cm2), using MCB to detect both GSH levels and activity in cells. In purified, intact, recently excreted and aged C. parvum oocysts, MCB fluorescence localized GSH in intact oocysts at several nuclear and cytoplasmic sporozoite foci (n = 2–6). The function of GSH as an endogenous free radical scavenger in UV-irradiated oocysts was not demonstrated. Other free radical scavengers are more active than GSH in UV-treated C. parvum oocysts, and MCB is unlikely to be useful as a surrogate for detecting UV damage in UV-treated Cryptosporidium oocysts. 2. UV disinfection induces apoptosis. The DNA intercalating dye YO-PRO1 (YP) was chosen to investigate the role of UV irradiation in inducing programmed cell death/apoptosis. YP binds strongly to nucleic acids and has been used to detect apoptosis in mammalian cells without interfering with cell viability. We focused our investigations on the usefulness of YP to detect DNA damage in UVtreated (40 mJ/cm2) C. parvum oocysts. YP was incorporated into sporozoite DNA of intact, irradiated oocysts (possibly apoptotic) which exhibited no apparent oocyst wall damage. Dexamethasone- or etoposide-induced apoptosis was localized at higher levels, compared with controls; however, control oocysts did not exclude YP entirely. No significant differences were detected between irradiated and drug-treated oocysts, compared with controls by fluorescence microscopy, but differences were apparent by FACS. Despite the increase in YP-positive oocysts by FACS, YP is unlikely to provide a reliable estimate of the possible apoptotic changes that can occur in irradiated oocysts or sporozoites. 3. UV disinfection causes damage to DNA, which is detectable using fluorogenic DNA reporters. The major DNA lesion caused by UV, the cyclobutyl

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pyrimidine dimer (TD), is responsible for UV-induced cytotoxicity and mutagenicity. It inhibits normal replication and transcription of DNA and results in the inactivation of cells. We used an antibody raised against TDs (α-TD) to identify changes induced by UV light in C. parvum oocysts, and validated its nuclear location by co-localization with the nuclear fluorogen, DAPI. To improve α-TD antibody labelling within irradiated C. parvum oocysts, a freeze-thawing (five cycles) procedure was developed. No α-TD localization was seen in non-irradiated oocysts. Replacing FITC-C-mAb with Texas Red (TR)-C-mAb and using the combination of α-TD-Ab, TR-C-mAb and DAPI, DNA damage to C. parvum and C. hominis sporozoites within intact oocysts could be detected at a lower limit of 10 mJ/cm2, but not at 4 mJ/cm2. Currently, the combination of α-TD-Ab, TR-CmAb and DAPI can be used to detect damage in nuclei of oocysts of C. parvum and C. hominis exposed to UV light (range 10–40 mJ/cm2). We conclude that an immunofluorescence assay using α-TD antibodies, which, for C. parvum, has been validated against a neonatal mouse infectivity assay, is suitable for detecting thymine dimers in air-dried oocysts and air-dried sporozoites of C. parvum and C. hominis oocysts, and that the α-dsDNA antibody is a good candidate for a positive control for the assay.

Acknowledgements This work was funded by the Environmental and Rural Affairs Department, Agricultural and Biological Research Group, Scottish Executive, Scotland, UK. We thank Dr D. Reid, Drinking Water Quality Unit, Scottish Executive, Edinburgh, for managing the project.

References Al-Adhami, B.H., Nichols, R.A.B., Kusel, J.R., O’Grady, J. and Smith, H.V. (2006) Cryptosporidium parvum sporozoites contain glutathione. Parasitology 133, 555–563. Al-Adhami, B.H., Nichols, R.A.B., 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. Applied and Environmental Microbiology 73, 947–955. Anderson, M.E. (1998) Glutathione: an overview of biosynthesis and modulation. Chemico-Biological Interactions 24, 1–14. Bukhari, Z. and LeChevallier, M. (2003) Assessing UV reactor performance for treatment of finished water. Water Science and Technology 47, 179–184. Bukhari, Z., Hargy, T.M., Bolton, J.R., Dussert, B. and Clancy, J.L. (1999) Mediumpressure UV for oocyst inactivation. Journal of the American Water Works Association 91, 86–94. Cacciò, S.M., Thompson, R.C.A., McLauchlin, J. and Smith, H.V. (2005) Unravelling Cryptosporidium and Giardia epidemiology. Trends in Parasitology 21, 430–437. Campbell, A.T., Robertson, L.J. and Smith, H.V. (1992) Viability of Cryptosporidium parvum oocysts: correlation of in vitro excystation with inclusion or exclusion of fluorogenic vital dyes. Applied and Experimental Microbiology 58, 3488–3493.

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Campbell, A.T., Robertson, L.J., Snowball, M.R. and Smith, H.V. (1995) Inactivation of oocysts of Cryptosporidium parvum by ultraviolet irradiation. Water Research 29, 2583–2586. Cheng, K.C., Cahill, D.S., Kasai, H., Nishimura, S. and Leob, L.A. (1992) 8-Hydroxyguanine, an abundant form of oxidative DNA damage, causes G-T and A-T substitution. Journal of Biological Chemistry 267, 166–172. Clancy, J.L., Bukhari, Z., McCuin, R., Clancy, T.P., Marshall, M.M., Korich, D.G., Fricker, C.R., Sykes, N., Smith, H.V., O’Grady, J.E., Rosen, J.P., Sobrinho, J. and Schaefer, F.W., III (2000) Cryptosporidium: Viability and Infectivity Methods. American Water Works Association Research Foundation / American Water Works Association / UK Drinking Water Inspectorate, pp.137 Cohen, G.M., Sun, X.M., Fearnhead, H., MacFarlane, M., Brown, D.G. Snowden, R.T. and Dinsdale, D. (1994) Formation of large weight fragments of DNA is a key committed step of apoptosis in thymocytes. Journal of Immunology 153, 507–516. Cohen, J.J. (1993) Apoptosis. Immunology Today 14, 126–130. 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 Research 35, 1387–1398. Drescher, A.C., Greene, D.M. and Gadgil, A.J. (2001) Cryptosporidium inactivation by low-pressure UV in a water disinfection device. Journal of Environmental Health 64, 31–35. El-Adhami, W., Daly, S. and Stewart, P.R. (1994) Biochemical studies on the lethal effects of solar and artificial ultraviolet radiation on Staphylococcus aureus. Archives of Microbiology 161, 82–87. Fayer, R., Morgan, U. and Upton, S.J. (2000) Epidemiology of Cryptosporidium: transmission, detection and identification. International Journal for Parasitology 30, 1305–1322. Fischer-Nielsen, A., Jeding, I.B. and Loft, S. (1994) Radiation-induced formation of 8-hydroxy-2-deoxyguanosine and its prevention by scavengers. Carcinogenesis 15, 1609–1612. Gilbert, M. and Knox, S. (1997) Influence of Bcl-2 overexpression on Na+/K+-ATPase pump activity: correlation with radiation-induced programmed cell death. Journal of Cellular Physiology 171, 299–304. Girdwood, R.W.A. and Smith, H.V. (1999) Cryptosporidium. In: Robinson, R., Batt, C. and Patel, P. (eds) Encyclopaedia of Food Microbiology, Vol. 1. Academic Press, London and New York, pp. 487–497. Haugland, R.P. (2005) Probes for cell adhesion, chemotaxis, multidrug resistance and glutathione. In: Spence, M.T.Z. (ed.) The Handbook: A Guide to Fluorescent Probes and Labelling Technologies. Molecular Probes Inc., Oregon, USA, pp. 767–776. Hijnen, W.A.M., Beerendonk, E.F. and Medema, G.J. (2006) Inactivation credit of UV irradiation for viruses, bacteria and protozoan (oo)cysts in water: a review. Water Research 40, 3–22. Idziorek, T., Estaquier, J., De Bels, F. and Ameisen, J.-C. (1995) YOPRO-1 permits cytofluorometric analysis of programmed cell death (apoptosis) without interfering with cell viability. Journal of Immunological Methods 185, 249–258. Johnson, A.M., Linden, K., Ciociola, K.M., De Leon, R., Widmer, G. and Rochelle, P.A. (2005) UV inactivation of Cryptosporidium hominis as measured in cell culture. Applied and Environmental Microbiology 71, 2800–2802. Karanis, P., Kourenti, C. and Smith, H. (2007) Waterborne transmission of protozoan parasites: a worldwide review of outbreaks and lessons learnt. Journal of Water Health 5, 1–38.

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H.V. Smith et al. Kochevar, I.E. (1990) UV-induced protein alterations and lipid oxidation in erythrocyte membranes. Photochemistry and Photobiology 52, 795–800. Korich, D.G., Marshall, M.M., Smith, H.V., O’Grady, J., Bukhari, Z., Fricker, C.R., Rosen, J.P. and Clancy, J.L. (2000) Inter-laboratory comparison of the CD-1 neonatal mouse logistic dose-response model for Cryptosporidium parvum oocysts. Journal of Eukaryotic Microbiology 47, 294–298. Kumar, A., Tyagi, M.B. and Jha, P.N. (2004) Evidences showing ultraviolet-B radiation-induced damage of DNA in cyanobacteria and its detection by PCR assay. Biochemical and Biophysical Research Communications, 318, 1025–1030. Lee, J.E., Park, J.C., Hwang, Y.S., Kim, J.K., Kim, J.G. and Sub, H. (2001) Characterization of UV-irradiated dense/porous collagen membranes: morphology, enzymatic degradation, and mechanical properties. Yonsei Medical Journal 42, 172–179. Lorenzo-Lorenzo, M.J., Ares-Mazas, M.E., Villacorta-Martinez de Maturana, I. and DuranOreiro, D. (1993) Effect of ultraviolet disinfection of drinking water on the viability of Cryptosporidium parvum oocysts. Journal of Parasitology 79, 67–70. Meister, A. and Anderson, M.E. (1983) Glutathione. Annual Review of Biochemistry 52, 711–760. Mitchell, D.L. (1988) The relative cytotoxicity of (6-4) photoproducts and cyclobutane dimers in mammalian cells. Photochemistry and Photobiology 48, 51–57. 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. Applied and Environmental Microbiology 68, 5387–5393. Nejedly, K., Kittner, R., Pospisilova, S. and Kypr, J. (2001a) Crosslinking of the complementary strands of DNA by UV light: dependence on the oligonucleotide composition of the UV irradiated DNA. Biochimica et Biophysica Acta 1517, 365–75. Nejedly, K., Kittner, R. and Kypr, J. (2001b) Genomic DNA regions whose complementary strands are prone to UV light-induced crosslinking. Archives of Biochemistry and Biophysics 388, 216–224. Nichols, R.A.B. and Smith, H.V. (2004) Optimisation of DNA extraction and molecular detection of Cryptosporidium parvum oocysts in natural mineral water sources. Journal of Food Protection 67, 524–532. O’Grady, J.E. and Smith, H.V. (2002) Methods for determining the viability and infectivity of Cryptosporidium oocysts and Giardia cysts. In: Ziglio, G. and Palumbo, F. (eds) Detection Methods for Algae, Protozoa and Helminths. John Wiley and Sons, Chichester, UK, pp. 193–220. Oguma, K., Katayama, H., Mitani, H., Morita, S., Hirata, T. and Ohgaki, S. (2001) Determination of pyrimidine dimers in Escherichia coli and Cryptosporidium parvum during UV light inactivation, photoreactivation, and dark repair. Applied and Environmental Microbiology 67, 4630–4637. Plantin-Carrenard, E., Bringuier, A., Derappe, C., Pichon, J., Guillot, R., Bernard, M., Foglietti, M.J., Feldmann, G., Aubery, M. and Braut-Boucher, F. (2003) A fluorescence microplate assay using Yopro-1 to measure apoptosis: application to HL60 cells subjected to oxidative stress. Cell Biology and Toxicology 19, 121–133. Robertson, L.J., Campbell, A.T. and Smith, H.V. (1993) In vitro excystation of Cryptosporidium parvum. Parasitology 106, 13–29. 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 repair genes. Journal of Eukaryotic Microbiology 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 in Parasitology 21, 81–87.

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Roza, L., De Gruijl, F.R., Bergen Henegouwen, J.B.A., Guikers, K., Van Weelden, H., Van Der Schans, G.P. and Baan, R.A. (1991) Detection of photorepair of UV-induced thymine dimers in human epidermis by immunofluorescence microscopy. Journal of Investigative Dermatology 96, 903–907. Shin, G.-A., Linden, K.G., Arrowood, M.J. and Sobsey, M.D. (2001) Low-pressure UV inactivation and DNA repair potential of Cryptosporidium parvum oocysts. Applied and Environmental Microbiology 67, 3029–3032. Slifko, T.R., Huffman, D.E. and Rose, J.B. (1999) A most-probable-number assay for enumeration of infectious Cryptosporidium parvum oocysts. Applied and Environmental Microbiology 65, 3936–3941. Slifko, T.R., Smith, H.V. and Rose, J.B. (2000) Emerging parasite zoonoses associated with food and water. International Journal for Parasitology 30, 1379–1393. Smith, H.V. and Rose, J.B. (1990) Waterborne cryptosporidiosis. Parasitology Today 6, 8–12. Smith, H.V. and Rose, J.B. (1998).Waterborne cryptosporidiosis: current status. Parasitology Today 14, 14–22. Smith, H.V., Robertson, L.J. and Ongerth, J.E. (1995) Cryptosporidiosis and giardiasis: the impact of waterborne transmission. Aqua – Journal of Water Supply: Research and Technology 44, 258–274. Smith, H.V., Cacciò, S.M., Cook, N., Nichols, R.A.B. and Tait, A. (2007) Cryptosporidium and Giardia as foodborne zoonoses. Veterinary Parasitology 149, 29–40. Takeshita, K., Shibato, J., Sameshima, T., Fukunaga, S., Isobe, S., Arihara, K. and Itoh, M. (2003) Damage of yeast cells induced by pulsed light irradiation. International Journal of Food Microbiology 85, 151–158. Wang, Y.-C., Lee, P.-J., Shih, C.-M., Chen, H.-Y., Lee, C.-C., Chang, Y.-Y., Hsu, Y.-T., Liang, Y.-J., Wang, L.-Y., Han, W.-H. and Wang, I.-C. (2003) Damage formation and repair efficiency in the p53 gene of cell lines and blood lymphocytes assayed by multiplex long quantitative polymerase chain reaction. Analytical Biochemistry 319, 206–215. Will, O., Mahler, H., Arrigo, A. and Epe, B. (1999) Influence of glutathione levels and heat-shock on the steady-state levels of oxidative DNA base modifications in mammalian cells. Carcinogenesis 20, 333–337. Xiao, L., Fayer, R., Ryan, U. and Upton, S.J. (2004) Cryptosporidium taxonomy: recent advances and implications for public health. Clinical Microbiological Reviews 17, 72–97.

16

Effect of Environmental and Conventional Water Treatment Processes on Waterborne Cryptosporidium Oocysts

B. KING, A. KEEGAN, C. SAINT AND P. MONIS Australian Water Quality Centre, Salisbury, Australia

Abstract Cryptosporidium oocysts are prevalent in surface waters as a result of anthroponotic activity and native animal faecal contamination. A sound understanding of the impact of environmental and water treatment processes on the survival of oocysts is required to properly risk-assess the threat posed to public health by the presence of oocysts in water storages. This chapter provides an overview of recently completed work studying the impact of temperature, predation, sunlight and conventional water treatment processes on oocyst survival in water.

Background Cryptosporidium oocysts are frequently found in surface waters (Smith and Rose, 1990; Rose et al., 1997) and are extremely resistant to chlorine and monochloramine at the concentrations used to disinfect water for potable use (Korich et al., 1990; Finch et al., 1993). In addition to being resistant to commonly used disinfectants, it is generally thought that oocysts can persist for several months or more in the aquatic environment (Robertson et al., 1992; Johnson et al., 1997; Medema et al., 1997). These factors, combined with the relatively low infectious dose demonstrated for some isolates of Cryptosporidium parvum (Messner et al., 2001), mean that Cryptosporidium represents a challenge to water utilities responsible for the provision of safe drinking water to the public. Water utilities require accurate data on the fate and transport of oocysts in water and their response to water treatment processes, in order to effectively riskassess and risk-manage the threat posed by the presence of oocysts in source waters. A number of studies have examined oocyst survival, using techniques such as in vitro excystation or vital dye staining (Robertson et al., 1992; Chauret et al., 1998), but such methods are only indicators of viability and are known to overestimate infectivity (Black et al., 1996; Bukhari et al., 2000). As a consequence, 198

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there is a lack of knowledge regarding the inactivation of oocysts in environmental waters, particularly with regard to the effect of biotic and abiotic processes on the retention of oocyst infectivity.

Objectives In order to address knowledge gaps with respect to the survival of oocysts in response to environmental and water treatment conditions, studies were conducted using a cell culture infectivity model (Keegan et al., 2003) to examine the response of oocysts in water to: ● ● ● ● ● ●

Temperature. Microbial activity. Natural light. Chemical flocculation (aluminium sulphate). Dissolved air flotation. Disinfection (chlorination, chloramination).

Approach Details of the methodologies used for the temperature, sunlight and treatment inactivation studies have been described in detail elsewhere (Keegan et al., 2003; King et al., 2005, 2007). Fresh or aged oocysts of Cryptosporidium parvum (Swiss cattle C26) that had been passaged through mice were used for all experiments, and infectivity was assessed using a cell culture infectivity/Taqman PCR assay (CC-PCR) (Keegan et al., 2003). A schematic diagram of the assay is presented in Fig. 16.1. For environmental survival experiments, microcosms containing fresh oocysts and either reagent-grade water, tap water or reservoir water were established in either sterile polycarbonate chambers (for temperature experiments) or methylacrylate chambers (for sunlight inactivation experiments). Oocyst infectivity was measured in triplicate using 10,000 oocysts, and log inactivation was calculated by comparing quantitative PCR results for the treated samples with the non-treatment controls (which represent maximum infectivity). Candidate predators of oocysts were enriched from Hope Valley reservoir water sediment by selecting for organisms with a food preference in the Cryptosporidium oocyst size-range, using cysts of the amoeba Rosculus spp. and baker’s yeast (Saccharomyces cerevisiae). Organisms were collected using micromanipulation and placed in an individual well of a 24-well Cellstar tissue culture plate (Greiner Bio-one, Frickenhausen, Germany) containing 1 ml of autoclaved Hope Valley reservoir water, supplemented with yeast dissolved in Ringer’s solution. Oocysts were pre-stained with a fluorescent antibody (AusFlow Cry 104; BTF, Sydney, Australia) and 4′,6-diamidino-2-phenylindole (DAPI) prior to feeding experiments. Feeding experiments were conducted using 8-well Lab-Tek II Chamber Slides (Nalgene Nunc International, Naperville, Illinois) for amoebae or 1.5 ml microcentrifuge tubes for other organisms in a total volume of 150 µl

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Treat to promote release of sporozoites, apply oocysts to cell monolayer

Incubate 48 hours to allow establishment of infection

Wash monolayer, extract DNA from monolayer, conduct quantitative real-time PCR

Analyse data, compare treatments versus untreated controls

Fig. 16.1. Overview of a Cryptosporidium infectivity assay that combines cell culture and real-time PCR.

of sterile reservoir water containing 5 × 104 oocysts per chamber. Samples were fixed by the addition 150 µl of sodium acetate acetic acid formalin (SAF). Organisms were viewed using an Olympus BX60 system microscope (Olympus, Oakleigh, Australia) and images were captured using an Olympus DP50 digital camera. The parameters used for conventional 2 l jar testing (to assess the effect of coagulation on oocyst infectivity) are listed in Table 16.1. Between 4 × 104 and 1 × 105 oocysts were used per jar, with doses of alum ranging from 0 to 100 mg/l,

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Experimental jar test conditions.

Jar test parameters Oocyst mixing Flash mixing Slow mixing Settling Volume Raw water chemistry pH UV254 Temperature Turbidity Colour

5 min @ 200 rpm 1 min @ 200 rpm 14 min @ 20 rpm 15 min 2 l or 400 ml 8.13–8.40 0.148–0.195/cm 16.8 ± 1.2°C 2.95–3.25 NTU 21–24 HU

which is within the range of concentrations used at conventional water treatment plants in South Australia. For samples further treated by dissolved air flotation (DAF), a 15 min flotation step with 12% recycle at 70 psi was added to the end of the standard jar test procedure. For disinfection, contact times for chlorine and chloramine ranged from 300 to 1200 mg min/l.

Outcomes Temperature inactivation Replicate in vitro experiments were performed in reagent-grade MilliQ water and Hope Valley raw water (with or without sterilization) to assess oocyst removal and inactivation rates at temperatures ranging from 4°C to 25°C. Infectivity for each time point was measured using 10,000 oocysts in triplicate, with inactivation determined by comparison with the level of infectivity observed in the 4°C control at each time point. The inactivation of the 4°C sample over time was calculated using the 4°C sample for t = 0 as the reference for maximum infectivity. The rate of inactivation was related to the temperature, with more rapid inactivation observed at the higher temperatures (Fig. 16.2). Additional experiments were conducted to investigate the mechanism responsible for oocyst inactivation by temperature. Oocysts were stored in reagent-grade water at temperatures between 4°C and 37°C and measurements were made for infectivity and oocyst ATP concentrations to determine whether there was any effect on oocyst energy stores. Changes in ATP concentration closely corresponded with changes in infectivity, with a good correlation observed between these two parameters for all temperatures (Fig. 16.3).

Log inactivation

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0.5 0.0 -0.5 -1.0 -1.5 -2.0 -2.5 -3.0 -3.5 -4.0 -4.5

4 degrees 15 degrees 20 degrees 25 degrees

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0

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4 6 8 Time (weeks)

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0.5 0.0 -0.5 -1.0 -1.5 -2.0 -2.5 -3.0 -3.5 -4.0 -4.5 0

2

4 6 8 Time (weeks)

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0.5 0.0 -0.5 -1.0 -1.5 -2.0 -2.5 -3.0 -3.5 -4.0 -4.5 12

0

2

4 6 8 Time (weeks)

10

12

Fig. 16.2. Inactivation of Cryptosporidium oocysts in water at different temperatures.

Predation Monitoring of oocyst numbers during the initial temperature inactivation study revealed changes in oocyst numbers for some treatments. In the case of the reagent water and autoclaved Hope Valley reservoir water treatments, the oocyst counts remained constant throughout the incubation period for all four temperatures in all replicate experiments. Furthermore, the morphology of the oocysts remained consistent, with a regular shape, good staining with the fluorescent antibody and no clumping observed. However, for one of the replicate experiments conducted on Hope Valley raw reservoir water, a gradual decrease in oocyst numbers was observed for the 15°C treatment and more rapid decreases for both 20°C and 25°C over the 12-week incubation period (Fig. 16.4). Oocysts examined in these samples were often clumped together with variable fluorescent staining and deformations in shape (as many as 18 in an individual clump, data not shown). Oocysts not in clumps exhibited typical staining and morphology, and exhibited infectivity consistent with oocysts in sterile water for the same temperature exposure.

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1E-10

0.5

ATP concentration (M) 1E-9

2E-9 1.8E-9 1.6E-9

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1.4E-9

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1.2E-9

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1E-9

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-1.5 -4

6E-10

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0

20

40

60

80

100

120

140

160

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Time (hours) incubation at 37°C

Fig. 16.3. Comparison of oocyst inactivation measured using cell culture infectivity with ATP concentration, measured during incubation at 37°C. Inset: correlation of oocyst ATP concentration and log inactivation using the combined data from all temperature treatments (15, 20, 25, 30, 37°C) and time points.

Attempts to replicate the oocyst removal in further experiments were variable between replicates, but decreases in oocyst numbers were observed. Reservoir water filtered using an 0.8 micron filter did not exhibit any reduction in oocyst numbers. To further investigate the loss of oocysts, potential predators of oocysts were enriched from the reservoir water and tested to determine whether they could ingest oocysts. A range of ciliates, amoebae and rotifers were identified that could ingest antibody-labelled oocysts, as determined by fluorescent microscopy and DAPI staining, as well as a single platyhelminth and gastrotrich (Table 16.2). In the case of Paramecium, it appeared that oocysts were being digested in food vacuoles (Fig. 16.5). In the case of Blepharisma, it was not possible to view the FITC-labelled oocysts due to autofluorescence, but the oocysts were clearly visible by DAPI staining.

Solar inactivation Outdoor tank experiments and a cell culture infectivity assay were used to measure the effect of solar irradiation on C. parvum oocysts in water. Initial experiments

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70 4°C

60

15°C 50

20°C 25°C

40

4°C autoclaved 30

15°C autoclaved

20

20°C autoclaved 25°C autoclaved

10 0

2

4 6 8 Incubation time (weeks)

10

12

Fig. 16.4. Comparison of oocyst counts in Hope Valley reservoir water (±sterilization) incubated at different temperatures over time.

Table 16.2.

List of phagotrophs capable of ingesting Cryptosporidium oocysts.

Phagotroph group

Taxon

Ciliates

Paramecium Euplotes Blepharisma Oxytricha Holosticha Mayorella Microchlamys Thecamoeba Willaertia Cochliopodium Unidentified rhabdocoel Lepadella Unidentified bdelloid (×3 types) Dicranophorus Chaetonotus

Amoebae

Platyhelminth Rotifers

Gastrotrich

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(B)

(C)

20

Fig. 16.5. Paramecium sp. containing ingested oocysts of Cryptosporidium parvum. (A) Oocysts are clearly visible in the food vacuoles (arrow) when viewed under differential interference contrast (DIC) optics. (B) The sporozoite nuclei within oocysts (arrows) can be seen with DAPI staining. (C) Fluorescent antibody-stained oocysts are visible (small arrow), with oocysts with decreased staining or signs of degradation indicated by a larger arrow. Bar = 20 µm.

Table 16.3. index days. UV indexa 1 3 4 4 7 10 11 12

Inactivation of Cryptosporidium oocysts in tap water on different UV

S90 (kJ/m2)b

T90 (h)c

13,200 8,700 5,300 6,200 11,900 4,400 5,900 76

6.4 3.7 1.8 1.7 2.6 0.9 1.0 0.4

a UV

index = estimated dose at solar noon, 1 = 25 mW/m2 UV between 290 and 400 nm. = the insolation necessary to achieve a 90% reduction in cell culture infectivity. c T90 = the time necessary to achieve a 90% reduction in cell culture infectivity. b S90

used tap water and assessed solar irradiation on days with a different UV index. Days with a UV index of 10 or greater resulted in rapid inactivation (Table 16.3). Dark controls (samples wrapped in aluminium foil), kept under the same conditions as the light-exposed oocysts, exhibited no change in infectivity compared with oocysts stored at 4°C, demonstrating that temperature did not affect oocyst infectivity within the time frame of these experiments.

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Solar inactivation was next assessed for reservoir waters that varied in turbidity, colour and dissolved organic carbon (DOC) (Table 16.4). Waters with high DOC required higher doses of solar irradiation to achieve oocyst inactivation (Table 16.5). Long pass filter experiments were conducted to determine the biologically active components of solar radiation. The component responsible for the majority of inactivation was found to be UV-B, with a small amount of inactivation attributed to UV-A. Visible light had no measurable effect, as determined by the cell culture infectivity assay. The level of UV-induced DNA damage was assessed using an antibody directed against cyclobutane thymine dimers and also by quantitative sequence detection using real-time PCR. The amount of damage detected was considerably less than expected considering the level of inactivation observed when compared to similar levels of oocyst inactivation using UV-C (254 nm), suggesting that other mechanisms in addition to DNA damage are responsible for oocyst inactivation.

Water treatment Conventional water treatment processes remove Cryptosporidium oocysts through coagulation, flocculation, sedimentation and filtration. Little is known regarding the effect of these processes on oocyst infectivity or whether exposure Table 16.4. ments.

Characteristics of reservoir water used for solar inactivation experi-

Water type Bolivar tap Warragamba Prospect Lake Hope Valley Myponga

pH

Dissolved organic carbon (mg/l)

7.80 7.45 7.15 7.78 7.55

2.8 2.6 3.6 8.0 12.3

Turbidity (NTU)

Colour (HU)

0.238 2.94 2.26 7.42 4.12

2 29 47 38 77

Table 16.5. Inactivation of Cryptosporidium oocysts in reservoir waters on different UV index days. Winter: UV index 3 Water type Bolivar tap Warragamba Prospect Hope Valley Myponga

S90 (kJ/m2) 9,300 7,900 13,300 14,751 27,933

T90 (h) 3.2 2.6 4.5 4.88 10.75

Summer: UV index 10 S90(kJ/m2) 4,500 5,800 5,200 13,500 32,599

T90 (h) 0.85 1.1 1.0 2.5 6.5

Waterborne Cryptosporidium Oocysts

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to these processes makes oocysts more susceptible to standard disinfection. The effect of the water treatment chemical alum on Cryptosporidium oocyst infectivity was assessed using cell culture infectivity. Coagulation by doses of alum ranging from 40 mg/l to 100 mg/l had no effect on oocyst infectivity (either on oocysts left in suspension or oocysts recovered from flocs). Dissolved air flotation of oocysts following alum coagulation also had no effect on oocyst infectivity. There was no change in the sensitivity of oocysts to chlorine or monochloramine following alum coagulation. The effect of age and temperature on oocyst sensitivity to chlorine was assessed (up to 6 months at 4°C or 15°C). A small increase in oocyst sensitivity to chlorine (Ct = 1200 mg min/l), resulting in 0.5 0 1 log inactivation, was observed for oocysts that had been stored at either 4°C or 15°C for 20 weeks or more (Fig. 16.6).

Summary and Future Directions While environmentally tough and resistant to common disinfection chemicals, Cryptosporidium oocysts are not invincible and are susceptible to environmental stresses. The data produced from this work need to be incorporated into fate and transport models, as current models use inaccurate data for temperature inactivation, and solar inactivation data have either been absent from such models or derived from other organisms. This will not only allow prediction of the location of oocysts in a reservoir but may also allow some prediction of infectivity if environmental conditions (temperature, light, DOC) are known. Importantly we also

1 week 0

week 8

week 12

week 16

week 20

week 24

Log survival

0 -1 -2 -3 -4 -5 Incubation period No treatment 4

JT 4

JT+cl2

No treatment 15

JT 15

JT+cl2 15

Fig. 16.6. Inactivation of oocysts by chlorine as a function of oocyst age and exposure to different temperatures.

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identify the need for further research effort directed towards biotic inactivation and removal. Questions which need to be answered include: ● ● ●

Are excreted oocysts still infectious? How does predation affect oocyst resistance to common disinfectants? Can such oocysts still be efficiently recovered by standard immunomagnetic separation (IMS) concentration?

Such work will provide valuable information for determining the relative risks associated with Cryptosporidium oocysts in water.

Acknowledgements We acknowledge the financial support received from the Co-operative Research Centre for Water Quality and Treatment, Water Services Association Australia, Australian Water Quality Centre, and South Australian Water Corporation. We thank David Daminato, Stella Fanok, Bret Robinson, Daniel Hoefel and Kylie Harvey for their excellent technical expertise and fruitful discussions.

References Black, E.K., Finch, G.R., Taghi-Kilani,