A laboratory manual for the isolation, identification and characterization of avian pathogens [5 ed.] 978-0-9789163-2-9

Table of contents :
Diagnostic Principles. Frederic J. Hoerr
Salmonellosis. W. Douglas Waltman and Richard K. Gast
Colibacillosis. Margie D. Lee and LisaK. Nolan
Pasteurellosis, Avibacteriosis, Gallibacteriosis, Riemerellosis, and Pseudotuberculosis.
John R Glisson, Tirath S. Sandhu, and Charles L. Hofacre
Bordetellosis. Mark W. Jackwood
Infectious Coryza. Pat J. Blackall
Campylobacter in Poultry. Jaap A. Wagenaar and Wilma F. Jacobs- Reitsma
Spirochetosis. Stephen R Collett
Erysipelas. George L. Cooper and Arthur A. Bickford
Listerosis. George L. Cooper and Arthur A. Bickford
Staphylococcosis. Stephan G. Thayer and W. Douglas Waltman Streptococcosis and Enterococcosis. Stephan G. Thayer and W. Douglas Waltman
Clostridial Diseases. Stephan G. Thayer and David A. Miller
Tuberculosis. Susan Sanchez and Richard M. Fulton
Mycoplasmosis. Stanley H. Kleven
Chlamydiosis. Arthur A. Andersen and Daisy Vanrompay
Ornithobacteriosis. Richard P. Chin and Bruce R Charlton Mycoses and Mycotoxicoses. RD. Wyatt
Adenovirus. Brian M. Adair and J. Brian McFerran
Hemorrhagic Enteritis of Turkeys Marble Spleen Disease of Pheasants. F. William Pierson and Scott D. Fitzgerald Infectious Laryngotracheitis. Deoki N. Tripathy and Maricarmen Garcia Marek’s Disease. Patricia S. Wakenell and Jagdev M. Sharma
Duck Virus Enteritis. Peter R. Woolcock
Herpesviruses of Free-Living and Pet Birds. Erhard F. Kaleta
Pox. Deoki N. Tripathy and Willie M. Reed
Budgerigar Fledgling Disease and Other Avian Polyomavirus Infections. Branson W. Ritchie and Phil D. Lukert Psittacine Beak and Feather Disease. Branson W. Ritchie and Phil D. Lukert
Chicken Anemia Virus. M. Stewart McNulty and Daniel Todd
Avian Influenza. David E. Swayne, Dennis A. Senne and David L. Suarez
Newcastle Disease Virus and Other Avian Paramyxoviruses. Dennis J. Alexander and Dennis A. Senne
Avian Metapneumovirus. Richard E. Gough and Janice C. Pedersen
Infectious Bronchitis. Jack Gelb, Jr. and Mark W. Jackwood
Turkey Coronavirus. Mark W. Jackwood and James S. Guy
Enteric Viruses. Don Reynolds and Ching Ching Wu
Oncomaviruses, Leukosis/Sarcomas and Reticuloendotheliosis. Aly M. Fadly, Richard L. Witter, and Henry D. Hunt
Avian Encephalomyelitis. Louis van der Heide
Duck Hepatitis. Peter R Woolcock
Turkey Viral Hepatitis. Willie M. Reed
Viral Arthr1t1s/Tenosynov1t1s and Other Reovirus Infections. John K. Rosenberger and Erica Spackman
Arbovirus Infection. Eileen N. Ostlund and James E. Pearson
Infectious Bursal Disease. John K. Rosenberger, Y.M. Saif, and Daral J. Jackwood
Parvovirus of Waterfowl. Richard E. Gough
Cell-Culture Methods. Karel A. Schat and Holly S. Sellers
Virus Propagation in Embryonating Eggs. Dennis A. Senne
Virus Identification and Classification. Pedro Villegas and Ivan Alvarado
Titration of Biological Suspensions. Pedro Villegas
Serologic Procedures. Stephan G. Thayer and Charles W. Beard
Molecular Identification Procedures. Daral J. JackwoOd and Mark W. Jackwood
Antigen Detection Systems. Mary J. Pantin-Jackwood and Sandra S. Rosenberger
Appendix of Abbreviations and Acronyms used in the Text
Quick Reference Diagnostic Chart
Index

Citation preview

A LABORATORY MANUAL FOR THE

ISOLATION, IDENTIFICATION, AND CHARACTERIZATION OF AVIAN -

UH*

**

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«

Fifth Edition

THE AMERICAN ASSOCIATION OF AVIAN PATHOLOGISTS

A LABORATORY MANUAL FOR THE

ISOLATION, IDENTIFICATION AND CHARACTERIZATION OF AVIAN PATHOGENS

Fifth Edition

American Association of Avian Pathologists Editorial Committee Louise Dufour-Zavala, Editor-in-Chief David E. Swayne John R. Glisson Janies E. Pearson Willie M. Reed Mark W. Jackwood Peter R. Woolcock

Copies Available from: American Association of Avian Pathologists 953 College Station Road Athens, GA 30602-4875

Frederic J. Hoerr

9 1

3

4

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5 1-------- 1 8

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Cover: 1. Direct fluorescent antibody test on primary chicken embryo kidney cells showing syncytia formed by ILT virus in primary chick embryo kidney cells. (Rey Resurreccion, GPLN, Oakwood, GA) 2. Ninety well serum plate prepared for ELISA testing for the detection of AE antibodies. (Len Chappell, GPLN, Oakwood, GA). 3. Bacterial culture on a TSI slant. (Doug Waltman, GPLN, Oakwood, GA). 4. Reverse transcriptase-polymerase chain reaction (RT-PCR) and restriction fragment length polymorphism (RFLP) analysis if the spike (SI) glycoprotein gene of the Arkansas strain of infectious bronchitis virus (IBV). Lanes 1 and 2 are molecular weight markers. Lane 3 is the IBV amplicon digested with BstYI, lane 4 is the IBV amplicon digested with Haein, and lane 5 is the IBV amplicon digested with XcmI. (Mark Jackwood, PDRC, Athens, GA). 5. Ten-fold dilution series of avian influenza virus RNA run with the USDA type A influenza M gene real-time RT-PCR test on the Applied Biosystems 7500 FAST system. (Erica Spackman, SEPRL, Athens, GA). 6. Microphotograph of Mycoplasma gallisepticum colonies on agar (35x) (Stan Kleven, PDRC, Athens, GA). 7. Bio Merieux API 20 system for bacteria identification using a series of biochemical tests. (Doug Waltman, GPLN, Oakwood, GA). 8. Embryonated egg candling to locate the chorio-allantoic sac for inoculation of a virus isolation sample. (Rey Resurreccion, GPLN, Oakwood, GA). 9. Wet mount of Aspergillus sp. conidiofores after culture in Sabouraud's dextrose agar (100X). The isolate was obtained from a case of systemic aspergillosis in 9-wk-old broiler breeder pullets. (Guillermo Zavala, PDRC, Athens, GA). Pictures 2,3,7,8 and Page design by HMM Photography (Heidi Migalla).

Copyright © 1975, 1980, 1989,1998,2008 by American Association of Avian Pathologists, Inc. Athens, Georgia

All rights reserved Copyright is not claimed for Chapters 22, 36 Chapters 16,29, 35,40,44, 49: US Government copyrighted, user rights reserved, printed by permission Chapters 30, 31: (British) Crown copyrighted and (British) Crown user rights reserved, printed by permission

Chapter 6: (Australian) Crown copyrighted and (Australian) Crown user rights reserved, printed by permission Mention of a trademark or proprietary product does not constitute a guarantee or warranty of the product by the author, the supporting institutions or the American Association of Avian Pathologists and does not imply its approval to the exclusion of other products that also may be suitable

Fifth edition 2008 Previously entitled Isolation and Identification ofAvian Pathogens Library of Congress Catalog Card Number: 2008924452

ISBN 978-0-9789163-2-9

All rights reserved. No part of this publication may be reproduced, stored in a retrieval system, or transmitted, in any form or by any means, electronic, mechanical, photocopying, recording, or otherwise, without the prior written permission of the copyright owner. Printed in the United States of America

10

987654321

Printed by OmniPress, Inc., Madison, Wisconsin 53704

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A Laboratory Manual for the Isolation, Identification and Characterization of Avian Pathogens

TABLE OF CONTENTS Diagnostic Principles. Frederic J. Hoerr...................................................................................................................................1 Salmonellosis. W. Douglas Waltman and Richard K. Gast.................................................................................................... 3 Colibacillosis. Margie D. Lee and Lisa K. Nolan..................................................................................................................10 Pasteurellosis, Avibacteriosis, Gallibacteriosis, Riemerellosis, and Pseudotuberculosis. John R. Glisson, Tirath S. Sandhu, and Charles L. Hofacre....................................................................................................12 5. Bordetellosis. Mark W. Jackwood........................................................................................................................................ 20 6. Infectious Coryza. Pat J. Blackall........................................................................................................................................ 22 7. Campylobacter in Poultry. Jaap A. Wagenaar and Wilma F. Jacobs- Reitsma..................................................................... 27 8. Spirochetosis. Stephen R. Collett........................................................................................................................................... 31 9. Erysipelas. George L. Cooper and Arthur A. Bickford........................................................................................................ 36 10. Listerosis. George L. Cooper and Arthur A. Bickford........................................................................................................ 39 11. Staphylococcosis. Stephan G. Thayer and W. Douglas Waltman.........................................................................................42 12. Streptococcosis and Enterococcosis. Stephan G. Thayer and W. Douglas Waltman............................................................ 44 13. Clostridial Diseases. Stephan G. Thayer and David A. Miller............................................................................................. 47 14. Tuberculosis. Susan Sanchez and Richard M. Fulton.......................................................................................................... 53 15. Mycoplasmosis. Stanley H. Kleven......................................................................................................................................59 16. Chlamydiosis. Arthur A. Andersen and Daisy Vanrompay................................................................................................. 65 17. Omithobacteriosis. Richard P. Chin and Bruce R. Charlton................................................................................................ 75 18. Mycoses and Mycotoxicoses. R.D. Wyatt............................................................................................................................ 77 19. Adenovirus. Brian M. Adair and J. Brian McFerran.............................................................................................................84 20. Hemorrhagic Enteritis of Turkeys Marble Spleen Disease of Pheasants. F. William Pierson and Sctott D. Fitzgerald....... 90 21. Infectious Laryngotracheitis. Deoki N. Tripathy and Maricarmen Garcia........................................................................... 94 22. Marek’s Disease. Patricia S. Wakenell and Jagdev M. Sharma........................................................................................... 99 23. Duck Virus Enteritis. Peter R. Woolcock............................................................................................................................ 106 24. Herpesviruses of Free-Living and Pet Birds. Erhard F. Kaleta........................................................................................... 110 25. Pox. Deoki N. Tripathy and Willie M. Reed....................................................................................................................... 116 26. Budgerigar Fledgling Disease and Other Avian Polyomavirus Infections. Branson W. Ritchie and Phil D. Lukert........ 120 27. Psittacine Beak and Feather Disease. Branson W. Ritchie and Phil D. Lukert................................................................... 122 28. Chicken Anemia Virus. M. Stewart McNulty and Daniel Todd......................................................................................... 124 29. Avian Influenza. David E. Swayne, Dennis A. Senne and David L. Suarez....................................................................... 128 30. Newcastle Disease Virus and Other Avian Paramyxoviruses. Dennis J. Alexander and Dennis A. Senne.........................135 31. Avian Metapneumovirus. Richard E. Gough and Janice C. Pedersen................................................................................. 142 32. Infectious Bronchitis. Jack Gelb, Jr. and Mark W. Jackwood............................................................................................. 146 33. Turkey Coronavirus. Mark W. Jackwood and James S. Guy.............................................................................................. 150 34. Enteric Viruses. Don Reynolds and Ching Ching Wu.................................................................................................... 153 35. Oncornaviruses, Leukosis/Sarcomas and Reticuloendotheliosis. Aly M. Fadly, Richard L. Witter, and Henry D. Hunt.. 164 36. Avian Encephalomyelitis. Louis van der Heide................................................................................................................... 173 37. Duck Hepatitis. Peter R. Woolcock............................................................................................................................ 175 38. Turkey Viral Hepatitis. Willie M. Reed.............................................................................................................................. 179 39. Viral Arthritis/Tenosynovitis and Other Reovirus Infections. John K. Rosenberger and Erica Spackman.........................181 40. Arbovirus Infection. Eileen N. Ostlund and James E. Pearson........................................................................................... 184 41. Infectious Bursal Disease. John K. Rosenberger, Y.M. Saif, and Daral J. Jackwood......................................................... 188 42. Parvovirus of Waterfowl. Richard E. Gough...................................................................................................................... 191 43. Cell-Culture Methods. Karel A. Schat and Holly S. Sellers................................................................................................ 195 44. Virus Propagation in Embryonating Eggs. Dennis A. Senne..............................................................................................204 45. Virus Identification and Classification. Pedro Villegas and Ivan Alvarado........................................................................209 46. Titration of Biological Suspensions. Pedro Villegas........................................................................................................... 217 47. Serologic Procedures. Stephan G. Thayer and Charles W. Beard.......................................................................................222 48. Molecular Identification Procedures. Daral J. Jackwood and Mark W. Jackwood..............................................................230 49. Antigen Detection Systems. Mary J. Pantin-Jackwood and Sandra S. Rosenberger.......................................................... 233 Appendix of Abbreviations and Acronyms used in the Text...................................................................................................... 241 Quick Reference Diagnostic Chart.............................................................................................................................................. 244 Index............................................................................................................................................................................................ 247

1. 2. 3. 4.

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CONTRIBUTING AUTHORS Brian Adair Veterinary Sciences Division Agri-Food and Biosciences Institute Stoney Road Stormont Belfast BT4 3SD Northern Ireland E-mail: [email protected] FAX: +44(0)2890525773

Richard P. Chin California Veterinary Diagnostic Laboratory System Fresno Branch School of Veterinary Medicine University of California - Davis 2789 S. Orange Ave. Fresno, California 93725 E-mail: [email protected] FAX: (559) 485-8097

Dennis J. Alexander Central Veterinaiy Laboratory (Weybridge) New Haw, Addlestone Surrey KT15 3NB United Kingdom E-mail: [email protected] FAX: +44-1932-357856

Stephen R. Collett Department of Population Health College of Veterinary Medicine The University of Georgia 953 College Stations Road Athens, Georgia 30602-4875 E-mail: [email protected] FAX: (706) 542-5630

Ivan R. Alvarado 120 Spring Lake Pointe Athens, GA 30605 E-mail: [email protected] FAX: (404) 506-9013

George L. Cooper School of Veterinary Medicine University of California-Davis Calif. Vet. Diagnostic Lab. System Turlock Branch P.O. Box P Turlock, California 95381 E-mail: [email protected] . FAX: (209) 667-4261

Dr. Arthur A. Andersen USDA-ARS Former Address: National Animal Disease Center P.O.Box 70, Ames, IA 50010 Email: [email protected]

Charles W. Beard 130 Red Fox Run Athens, GA 30605 E-mail: [email protected] FAX: (706) 548-6410

Aly M. Fadly USDA- Agricultural Research Service Avian Disease and Oncology Laboratory (ADOL) 3606 East Mount Hope Road East Lansing, Michigan 48823 E-mail: [email protected] FAX: (517) 337-6776

Arthur A. Bickford University of California-Davis California Veterinaiy Diagnostic Laboratory System Turlock Branch 1650 Simon Drive Turlock, California 95382 E-mail: [email protected] FAX: (209) 632-4258

Scott D. Fitzgerald Diagnostic Center for Population and Animal Health College of Veterinary Medicine Michigan State University PO Box 30076 Lansing, Michigan 48909-7576 E-mail: [email protected] FAX: (517)355-2152

Pat J. Blackall Animal Research Institute Locked Mail Bag No. 4 Moorooka, Queensland 4105 Australia E-mail: [email protected] FAX: +61-7-33629-429

Richard M. Fulton Diagnostic Center for Population and Animal Health Michigan State University PO Box 30076 Lansing, Michigan 48909 E-mail: [email protected] FAX: (517)355-2152

Bruce R. Charlton California Veterinary Diagnostic Laboratory System Turlock Branch School of Veterinary Medicine University of California - Davis 3327 Chicharra Way Coulterville, California 95311 E-mail: [email protected] FAX: (209) 667-4267

Maricarmen Garcia Department of Population Health College of Veterinary Medicine The University of Georgia 953 College Station Rd. Athens, Georgia 30602-4875 E-mail: [email protected] FAX: (706) 542-5630

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Richard K. Gast USDA, ARS Russell Research Center 950 College Station Road Athens, GA 30605 E-mail: [email protected] FAX: (706) 546-3035

Daral J. Jackwood Food Animal Health Research Program Ohio Agricultural Research and Development Center The Ohio State University 1680 Madison Ave Wooster, Ohio 44691 E-mail: [email protected] FAX: (330) 263-3760

Jack Gelb, Jr. Department of Animal and Food Sciences College of Agricultural Sciences 531 South College Avenue University of Delaware Newark, Delaware 19716-2150 E-mail: [email protected] FAX: (302)831-2822

Mark W. Jackwood Department of Population Health College of Veterinary Medicine The University of Georgia 953 College Station Road Athens, Georgia 30602-4875 E-mail: [email protected] FAX: (706) 542-5630

John R. Glisson Department of Population Health College of Veterinary Medicine The University of Georgia 953 College Station Rd. Athens, Georgia 30602-4875 E-mail: [email protected] FAX: (706) 542-5630

Wilma F. Jacobs-Reitsma RIKILT Institute of Food Safety Bomsesteeg 45 6708 PD Wageningen The Netherlands E-mail: [email protected] FAX: +31-317-417717

Richard E. Gough Central Veterinary Laboratory (Weybridge) New Haw, Addlestone Surrey KT15 3NB United Kingdom E-mail: [email protected] FAX: +44-1932-357856

Erhard F. Kaleta Institute for Avian and Reptile Medicine Justus-Liebig-University Frankfurter Strabe 91, D-35392 Giefien Germany E-mail: [email protected] FAX: +49-641-201548

Janies S. Guy North Carolina State University College of Veterinary Medicine Department of Population Health & Pathobiology 4700 Hillsborough Street Raleigh, North Carolina 27606 E-mail: Jim [email protected] FAX: (919)513-6464

Stanley H. Kleven Department of Population Health College of Veterinary Medicine University of Georgia 953 College Station Rd Athens, Georgia 30602-4875 E-mail: [email protected] FAX: (706) 542-5630

Frederic J. Hoerr Thomas Bishop Sparks State Diagnostic Laboratory P. O. Box 2209 Auburn, Alabama 36831-2209 E-mail: [email protected] FAX: (334) 8244-7206

Margie D. Lee Department of Population Health Poultry Diagnostic and Research Center The University of Georgia 953 College Station Road Athens, Georgia 30602-4875 E-mail: [email protected] FAX: (706) 542-5771

Charles Hofacre Department of Population Health College of Veterinary Medicine The University of Georgia 953 College Station Road Athens, Georgia 30602-4875 E-mail: [email protected] FAX: (706) 542-5630

Phil D. Lukert Department of Medical Microbiology College of Veterinary Medicine University of Georgia Box 207 Colbert, Georgia 30628 E-mail: [email protected] FAX: (706) 542-5771

Henry Hunt USDA- Agricultural Research Service Avian Disease and Oncology Laboratory (ADOL) 3606 East Mount Hope Road East Lansing, Michigan 48823 E-mail: [email protected] FAX: (517) 337-6776

J. Brian McFerran 19 Knocktem Gardens Belfast BT4 3LZ Northern Ireland FAX: +44-1232-658040

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M. Stewart McNulty Department of Agriculture Veterinary Sciences Division Stormont, Belfast BT4 3SD Northern Ireland FAX: +44-1232-525773 E-mail: m-mcnulty @utvintemet. com

Willie M. Reed Purdue University Dean, School of Veterinary Medicine Lynn Hall Room 1176 West Lafayette, Indiana 47907-2026 E-mail: [email protected] FAX: (765) 496-1261

David A. Miller National Veterinary Services Laboratories USDA-APHIS-VS 1800 Dayton Road Ames, Iowa 50010 E-mail: [email protected] FAX: (515) 239-8569

Donald L. Reynolds 2520 Veterinary Administration Iowa State University 1802 Elwood Drive Ames, Iowa 50011 E-mail: [email protected] FAX: (515) 294-8956

Lisa K, Nolan Dept, of Vet Microbiology and Preventive Medicine College of Veterinary Medicine 1802 Elwood Dr VMRI #2 Iowa State University Ames, LA 50011 E-mail: [email protected] FAX: (515) 233-2136

Branson W. Ritchie Department of Medical Microbiology College of Veterinary Medicine University of Georgia Athens, Georgia 30602 FAX: (706) 542-6460 Email: [email protected] John K. Rosenberger Aviserve, LLC Delaware Technology Park 1 Innovation Way, Suite 100 Newark, DE 19711 E-mail: [email protected] FAX: (302) 368-2975

Eileen N. Ostlund Head Equine Ovine Viruses Section Diagnostic Virology NVSL Ρ,Ο,Βοχ 844 Ames, IA 50010 E-mail: [email protected] FAX: (515) 663-7348

Sandra S. Rosenberger Aviserve, LLC Delaware Technology Park 1 Innovation Way, Suite 100 Newark, DE 19711 E-mail: [email protected] FAX: (302) 368-2975

Mary J. Pantin-Jackwood Southeast Poultry Research Laboratory USDA-ARS 934 College Station Road Athens, Georgia 30605 E-mail: [email protected] FAX: (706) 546-3161

Y. M. Saif Food Animal Health Research Program Ohio Agricultural Research and Development Center The Ohio State University 1680 Madison Avenue Wooster, Ohio 44691 E-mail: [email protected] FAX: (330) 263-3677

Janies E. Pearson 4016 Phoenix Ames, Iowa 50014 (515) 292-9435 E-mail: [email protected] Janice C. Pedersen Avian Section, Diagnostic Veterinary Services Laboratory NVSL 1437 270th Street Madrid, Iowa 50156 E-mail: [email protected] FAX: (515) 663-7348

Susan Sanchez Athens Diagnostic Laboratory 0103 Athens V.M. Diag Lab 501 D.E. Brooks Dr Athens, GA 30602 E-mail: [email protected] FAX: (706)542-5568

Frank W. Pierson Center for Molecular Medicine and Infectious Diseases Virginia-Maryland Reg. College of Vet. Medicine Virginia Polytechnic Institute and State University Blacksburg, Virginia 24061 E-mail: [email protected] FAX: (540) 231-3426

Tirath S. Sandhu Cornell University Duck Research Laboratory 37 Howell Place P.O. Box 427 Speonk, New York 11972 E-mail: [email protected]

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Stephen G. Thayer Department of Population Health College of Veterinary Medicine The University of Georgia 953 College Station Road Athens, Georgia 30602-4875 E-mail: [email protected] FAX: (706) 542-0252

Karel A. Schat Unit of Avian Health College of Veterinary Medicine Cornell University Ithaca, New York 14853 E-mail: [email protected] FAX: (607) 253-3384

Daniel Todd Dept of Agriculture and Rural Development from N. Ireland Veterinary Sciences Division Stormont, Belfast BT4 3SD UK E-mail: [email protected] Tel: +44 2890 525773

Holly Sellers Department of Population Health Poultry Diagnostic and Research Center The University of Georgia 953 College Station Road Athens, GA 30602-4875 E-mail: [email protected] FAX: (706)542-5630

Deoki N. Tripathy Department of Veterinary Pathobiology College of Veterinary Medicine University of Illinois 104 West McHenry Urbana, Illinois 61801 E-mail: [email protected] FAX: (217) 244-7421

Dennis A. Senne National Veterinary Services Laboratories Veterinary Services Animal and Plant Health Inspection Service United States Department of Agriculture 1800 Dayton Road Ames, Iowa 50010 E-mail: [email protected] FAX: (515) 663-7348

Daisy Vanrompay Ghent University Faculty of Bioscience engineering Department of molecular biotechnology Coupure Links 653 9000 Ghent, Belgium E-mail: [email protected] FAX: +32 09 2646219

Jagdev M. Sharma Department of Veterinary PathoBiology 258 Veterinary Science Building College of Veterinary Medicine University of Minnesota 1971 Commonwealth Avenue St. Paul, Minnesota 55108 E-mail: sharmOO 1 @umn.edu FAX: (612) 625-5203

Louis Van Der Heide Dept. Of Pathobiology U-89 PO Box 37 12 Yeomans Road Columbia, Connecticut 06237 E-mail: [email protected] FAX: (860) 486-2794

Erica Spackman USDA-Agriculture Research Service Southeast Poultry Research Laboratory 934 College Station Road Athens, GA 30605 E-mail: [email protected] FAX: (706)546-3161

Pedro Villegas Department of Population Health College of Veterinary Medicine University of Georgia 953 College Station Road Athens, Georgia 30602-4875 E-mail: [email protected] FAX: (706) 542-5630

David L. Suarez USDA-Agriculture Research Service Southeast Poultry Research Laboratory 934 College Station Road Athens, GA 30605 E-mail: [email protected] FAX: (706)546-3161

Jaap A. Wagenaar Department of Infectious Diseases and Immunology OIE Reference Laboratory for Campylobacteriosis and WHO Collaboration Centre for Campylobacter Faculty of Veterinary Medicine Utrecht University P.O. Box 80.165 3508 TD Utrecht The Netherlands E-mail: i [email protected] FAX: +31 (0)30-2533199

David E. Swayne USDA-Agricultural Research Service Southeast Poultry Research Laboratory 934 College Station Road Athens, Georgia 30605 E-mail: [email protected] FAX: (706) 546-3161

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Patricia S. Wakenell Department of Veterinary Medicine Population Health & Reproduction University of California Davis, California 95616 E-mail: pswakenell@ucdavis. edu FAX: (530) 752-5845

Peter R. Woolcock California Veterinary Diagnostic Laboratory System Fresno Branch School of Veterinary Medicine University of California - Davis 2789 South Orange Ave Fresno, California 93725 E-mail: [email protected] FAX: (559) 485-8097

W. Douglas Waltman Georgia Poultry Laboratory P.O. Box 20 4457 Oakwood Road Oakwood, Georgia 30566 E-mail: [email protected] FAX: (770) 535-1948

Ching-Ching Wu Purdue University Animal Disease Diagnostic Laboratory 406 South University Street West Lafayette, Indiana 47907-2065 E-mail: [email protected] FAX: (765) 497-1405

Richard L. Witter USDA- Agricultural Research Service Avian Disease and Oncology Laboratory 3606 East Mount Hope Road East Lansing, Michigan 48823 E-mail: [email protected] FAX: (517) 349-0817

Roger Wyatt 195 Edgewood Dr. SW Athens, GA 30606 (706) 548-2297 E-mail: [email protected]

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PREFACE This manual has its origins in the need for a book to codify standardized method for testing and evaluating poultry vaccines. The National Academy of Sciences sponsored the original publication entitled Methods for the Examination of Poultry Biologies, and completed two successive revisions. In 1975, the American Association of Avian Pathologists (AAAP) accepted responsibility for the publication, but because the need for standardizing testing of vaccines had been met, the scope and purpose of the manual was broadened to be a resource for laboratory procedures for the isolation and identification of disease-causing agents. The title was changed to Isolation and Identification of Avian Pathogens for the 1st edition and to A Laboratory Manual for the Isolation and Identification of Avian Pathogens for the 3rd edition. The manual was intended as a bench resource for daily use in the diagnostic laboratory.

With the 4th edition, the focus had shifted from a manual of procedures for isolation and identification of pathogens to a more encompassing manual for diagnosis of the disease and isolation and/or demonstration of the pathogen. This change was needed because many clinical specimens are presented as unknowns for determination of etiology and some pathogens are difficult to isolate, but can be demonstrated with newer molecular or immunologic techniques. With the 5th edition, the AAAP appointed Dr. Louise Dufour-Zavala as Editor-in-Chief. David Swayne served as advisor to the Editorin-Chief and section editor. John Glisson, Willie M. Reed, Mark W. Jackwood and James E. Pearson continued as section editors for their expertise in bacteriology, veterinary diagnostics, molecular biology, and virology. Peter Woolcock was newly appointed to support the virology section.

The 5th edition has a new title to reflect the molecular advances and modem testing procedures allowing us to characterize, type, speciate and serotype several avian pathogens. It also has a new chapter on Turkey Coronavirus. A color plate with tissue culture images is included. Improvements to individual chapters include updated terminology and information on molecular techniques. In the appendix section, we removed the appendix of sources and the appendix of reference antisera as they are now readily available on the Internet.

The editorial committee thanks all of the 67 contributors who prepared new chapters or revised existing chapters for the 5th edition. We also thank Crissie Boyd, Georgia Poultry Laboratory Network, for her tremendous clerical support in formatting the Manual. We thank Heidi Migalla, HMM photography, for the design of the cover and Aaron Nord at Omni Press for the printing services. Editorial Committee: Louise Dufour-Zavala, Editor-in-Chief David E. Swayne John R. Glisson Mark W. Jackwood James £. Pearson Willie M. Reed Peter Woolcock

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X

1 Diagnostic Principles Frederic J. Hoerr In avian diagnostic medicine the usual immediate challenge is making a definitive diagnosis for the presenting problem of morbidity or mortality. When the case is probed deeper, however, evidence may emerge of a concurrent or preceding infectious disease, management or nutritional problem, or other condition that contributes to the presenting problem. In food supply medicine involving a population of animals, the challenge is to evaluate one or more diagnoses for priority of response. In practice, the diagnostic results are considered along with animal welfare, production economics, and public health in the development of strategies for treatment, prevention, and control of the disease. Three factors influence the expression of an infectious disease: the virulence of the causative organism, the level of exposure or dose of the inoculum, and the susceptibility of the host. Within a poultry flock, each individual reacts according the net influence of these factors. The uniformity of exposure and resistance among the individuals in the flock will eventually influence flock performance and possibly food safety.

pneumovirus, infectious bronchitis, infectious laryngotracheitis, infectious bursal disease, chicken infectious anemia, reovirus, hemorrhagic enteritis virus, mycoplasmosis, infectious coryza, pasteurellosis, and others. For example, isolation of infectious bronchitis virus is an important first step in understanding a respiratory disease. If the virus is determined to be a new serotype, a significant change in vaccine virus serotype will be required to control the disease. If the isolated virus is indistinguishable from a vaccine strain administered at an earlier age, it may indicate the need for improvements in vaccine administration, or point to immunosuppression from infectious bursal disease, chicken infectious anemia, or other diseases. Some vaccine viruses are readily isolated for several days post­ administration. Extended periods of virus shedding in a flock may reflect variable immunity existing at the time of vaccine administration. For respiratory viruses, isolation of a vaccine virus for a prolonged period after administration can be indicative of socalled rolling reactions. In this situation, poor uniformity in flock immunity will result in some chicks being resistant to virus infection at the time of vaccine administration, then becoming susceptible during the end of the shedding period of a pen mate. The attenuated virus infection spreads from bird-to-bird resulting in vaccination reactions of prolonged duration. Attenuated vaccine viruses may re-acquire virulence characteristics during this process. Rolling reactions and lengthened periods of vaccine virus isolation also occur with uneven or ineffective vaccine administration technique. Vaccine virus shed from infected pen mates eventually infects susceptible chicks causing uneven and sometimes harsh reactions within the flock. Differentiating vaccine strains from wild­ type or field-challenge viruses or bacteria in the laboratory may require molecular genetic sequencing and analysis. The presence of co-pathogens can increase the severity of a viral disease such as infectious bronchitis or the respiratory form of Newcastle disease. Mycoplasma infections and elevated concentrations of ammonia and airborne dust increase the severity of respiratory viral disease. In these situations, the actual cause of death or economic loss in the form of condemnations at slaughter from respiratory compromise or septicemia caused by Escherichia coli. While the laboratory effort may focus on the viral infection, the cumulative effects of the co-pathogens must also take into consideration for effective treatment and control. This situation requires a comprehensive approach to diagnostics, including virology, bacteriology, serology, and pathology. Specific-pathogen-free sentinel birds are useful in identifying specific primary infectious pathogens. These are typically used when interference from overwhelming co-pathogens or vaccine strains obscures the isolation effort of primary pathogen. Sentinel birds can be placed in a flock for a specific time and then brought into the laboratory for pathogen isolation and identification procedures. Selective immunization of the sentinels focuses the susceptibility and thereby increases the efficiency of identifying the challenge strain. The relative significance of an isolate also depends on whether it has food safety or public health significance. For example, Salmonella, Escherichia, and Campylobacter can be pathogenic for poultry but even in the apparent absence of disease they can be significant contaminants on processed poultry or poultry products. Avian influenza virus has strain-variable virulence among different poultry species, and some strains have considerable public health significance.

HISTORICAL PERSPECTIVE

The very existence of this book makes a statement about the importance of infectious diseases that affect avian species. Detection and characterization of infectious pathogens have advanced substantially in recent decades, building on classical methods for cultivation, isolation, characterization, and immunological assay. Procedures based on the specific molecular genetic characteristics of disease agents are now extensively applied for the detection and characterization of infectious pathogens. Diagnostic technology today uses a variety of sensitive and specific methods that may not require the actual cultivation and isolation of an organism for a diagnosis. Isolation of an agent still has an important place in the investigative process. Isolates are important in assessing virulence and pathogenesis, and in the development of vaccines. Detection technologies offer significant gains and substantial advantages in the number of samples that can be tested in a short time, and in the overall efficiency of testing. Enzymelinked immunological detection (antigen capture ELISA) is used for viral and bacterial diseases. The polymerase chain reaction and nucleotide probes offer many approaches for identification of fragments of genetic code specific to a species or biotype of organism. A panel or multiplex of serological assays can reveal the spectrum of infectious agents that have infected an individual bird. Simultaneous statistical analysis of ELISA titers for various infectious pathogens helps to assess the frequency and distribution of the infection or vaccine-induced humoral immunity within the flock. Advances in diagnostic expertise that improve sensitivity and specificity, and reduce the time required for the test generally lead to improvements in disease prevention and control. This is essential to safeguarding poultry health in the ever larger size of poultry flocks under the care of food supply medicine.

SIGNIFICANCE OF IDENTIFIED PATHOGENS An isolated or detected disease agent should be evaluated for diagnostic significance relative to the case history, clinical disease, and lesions. Poultry in commercial flocks may experience sequential or simultaneous disease challenges, as well as exposure to agents in attenuated live vaccines. In a diagnostic investigation, both field challenge agents and attenuated live agents may be detected and require further characterization and differentiation. Examples of live vaccines include those for Newcastle disease

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microorganisms. Maintenance and calibration of equipment should include but not be limited to scales, precision pipettes, incubators, refrigerators, freezers, autoclaves, spectrophotometers and other visualization apparatuses, and electrophoresis equipment. Recording, documentation and review of laboratory procedures should be routine. Commercial test kits should be used according to manufacturer’s recommendations. Inventories of reagents and test kits should be managed with respect to expiration dates.

DISEASE REPORTING

The isolation and identification of certain pathogens carries the responsibility of reporting to state/provincial or federal regulatory officials, who in turn have responsibilities to report internationally to the World Organization for Animal Health (OIE) (1). The OIE website maintains a current list of internationally reportable diseases which includes Newcastle disease, turkey rhinotracheitis, any H5 or H7 avian influenza, Pullorum disease, and others. Reporting regulations vary among countries, states, and provinces, as do the definitions of reportable strains of an agent. Laboratory diagnosticians generally report to the next level of authority in their organization. Question about the responsibility to report a disease should be directed to the state or regional veterinary regulatory health official. Trade, regulatory and other legal issues may surround the isolation and accurate identification of certain pathogens, or serological evidence thereof. Erroneous reporting, misidentification or failure to identify an agent may carry serious consequences. In the United States, the USDA National Veterinary Services Laboratory is the agency that should be consulted about agent identification, and in other countries, the appropriate central reference laboratory.

LABORATORY SAFETY

The avian diagnostic laboratory today should operate under standard biosafety criteria (4). Although some avian procedures can be conducted at biosafety level 1, biosafety level 2 is desirable because some avian pathogens are agents of moderate potential hazard to personnel. Biosafety level 3 may be required for viruses and bacteria that may cause serious or potentially lethal disease as a result of exposure by the inhalation route. Salmonella, Campylobacter, Chlamydia, Erysipelas, Escherichia and Newcastle disease can infect persons causing clinical disease from mild conjunctivitis to systemic illness. Certain strains of highly pathogenic influenza virus may cause human fatalities. Fungal cultures carelessly handled can result in massive release of spores into the laboratory environment. Laboratory workers should be informed about these risks and trained in proper procedures for routine and specialized aspects of laboratory duties. Laboratories should be kept clean and orderly, and surfaces frequently cleaned and disinfected. Warning notices should be displayed by laboratories which contain hazardous substances or conditions. Biosafety cabinets of adequate rating should be maintained and fully functional. Eating, drinking, and smoking should be prohibited in laboratory work areas. Personal protective equipment including appropriate laboratory clothing is always in order, and respiratory masks and protective eye glasses should be worn when zoonotic pathogens are suspected.

QUALITY MANAGEMENT A quality management program to include standard operating procedures for the isolation and identification of infectious pathogens improves the overall consistency and quality of the laboratory effort. A quality management program evaluates laboratory procedures for the yield of relevant and timely data. It involves quality assessment by specifying performance parameters and setting limits for acceptable performance, and quality improvement by correcting problems and preventing their reoccurrence. Quality management helps to ensure that the laboratory information is accurate, reliable and reproducible, with the goal of eliminating or reducing test variation within and between laboratories. The elements of a quality program comprise written procedure manuals, record keeping, documentation, and retention, training, education and evaluation of personnel, proficiency testing, laboratoiy safety, and maintenance and monitoring of equipment. The International Standards Organization (ISO) is a developer and publisher of international standards for laboratoiy quality management (2). ISO is a non-governmental organizational network composed of the national standards institutes that currently represent 157 countries. ISO document 17025 is the main standard for diagnostic laboratories worldwide, including the USDA National Veterinary Services Laboratory. These standards are reflected in the essential requirements for accreditation by the American Association of Veterinary Laboratoiy Diagnosticians (AAVLD), which accredits publically-supported diagnostic laboratories in the United States and Canada (3). The AAVLD defines laboratoiy quality management to involve validation of test methods, among other criteria. Validation of a test requires ongoing documentation of internal or inter-laboratory performance using known reference standard(s) for the species and/or diagnostic specimen(s) of interest, and one or more of the following: the test is endorsed or published by reputable technical organization (including the American Association of Avian Pathologists Isolation and Identification of Avian Pathogens); published in a peer-reviewed journal with sufficient documentation to establish diagnostic performance and interpretation of results; or documentation of an internal or inter-laboratory comparison to an accepted methodology or protocol. In the avian diagnostic laboratory, quality management may include the utilization of check tests, reference sera, and reference strains of infectious agents, and maintenance of stock cultures of

STORAGE AND ARCHIVING OF ISOLATES A diagnostic laboratory has a wealth of information and valuable material pass through it. The agents isolated from commercial poultry represent a selection process involving large homogeneous animal populations under performance rearing conditions. These isolates have value for investigations of virulence and pathogenesis; epidemiology of emerging agents; evaluation of new drugs and vaccines, and potentially as new vaccines. Isolates from noncommercial poultry have value in comparative studies and in epidemiology. All viruses and most bacteria require ultracold (-70 F or lower) storage that can be achieved with specialty freezers or liquid nitrogen storage. Some bacteria and fungi can be stored in special media at room temperature. Specific information about storage and archiving of samples is provided in the text.

References 1. OIE: World Organization for Animal Health, http://www.oie.int/eng/maladies/en classification2007.htm?eld7. 2. International Organization for Standardization, http://www.iso.org/iso/home.htm. 3. American Association of Veterinary Laboratory Diagnosticians. AAVLD Essential Requirements for An Accredited Veterinary Medical Diagnostic Laboratory. http://web.memberclicks.com/mc/page.do?sitePageId=27915&orgl d=aavld. 4. Centers for Disease Control and Prevention. BMBL Section ΙΠ: Laboratoiy Biosafety Criteria. http: //www.cdc. gov/od/ohs/biosfty/bmbl4/bmbl4 s3 .htm.

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2 SALMONELLOSIS W. Douglas Waltman and Richard K. Gast

SUMMARY. Avian Salmonella infections are important as both a cause of clinical disease in poultry and as a source of food-bome transmission of disease to humans. Host-adapted salmonellae (5. pullorum and S. gallinarum) are responsible for severe systemic diseases, whereas numerous serotypes of non-host-adapted paratyphoid salmonellae are often carried subclinically by poultry and thereby may contaminate poultry products. Crowding, mal-nutrition, and other stressful conditions as well as unsanitary surroundings can exacerbate mortality and performance losses due to salmonellosis, especially in young birds. Agent Identification. Salmonella infections in poultry flocks are diagnosed principally by the isolation of the bacteria from clinical tissue samples, such as pooled internal organs and sections of the intestinal tract. Egg contents may also be cultured to detect S. enteritidis. Preliminary identification of infected or colonized flocks is often obtained by culturing poultry house or hatchery environmental samples, such as drag swabs, litter, dust, feed, and hatch residue. Clinical and environmental samples are generally subjected to selective enrichment, with nonselective preenrichment sometimes employed when salmonellae are present in very small numbers or are potentially injured. Delayed secondary enrichment often improves Salmonella recovery from many sample types. Samples are plated on selective agar and bacterial colonies are identified as Salmonella by further biochemical and serological testing. Serologic Detection in the Host. Serologic identification of infected poultry plays an important role in programs for controlling the spread of Salmonella in commercial flocks, especially in regard to S. pullorum and 5. gallinarum. Agglutination tests, particularly the rapid whole­ blood plate test, are used for verification of the Salmonella status of flocks participating in the National Poultry Improvement Plan.

measures be followed any time individuals go onto a farm or into a hatchery.

INTRODUCTION Avian Salmonella infections are important as both a cause of clinical disease in poultry and as a source of food-bome transmission of disease to humans. Host-adapted salmonellae are responsible for pullorum disease (S. pullorum) and fowl typhoid (S. gallinarum), which are severe systemic diseases that have become relatively rare in countries with testing and eradication programs (13). Infections with non-host-adapted (paratyphoid) salmonellae are common in all types of birds (9,10), although some serotypes are found predominantly in a fairly limited range of hosts (e.g., S. arizonae is isolated mostly from turkeys). Crowding, mal­ nutrition, and other stressful conditions as well as unsanitary surroundings can exacerbate mortality and performance losses due to salmonellosis. Paratyphoid salmonellae can also cause human illness, and food-bome Salmonella outbreaks can lead to severe economic losses to poultry producers as a result of regulatory actions, market restrictions, or reduced consumption of poultry products.

Clinical Cases Samples taken from the internal organs of the birds with systemic salmonellosis are typically free of competing bacteria. In such cases, isolation is usually relatively simple, and may only require direct plating on nonselective media. Liver, spleen, heart, heart blood, ovary or yolk sac, synovial fluid, eye, and brain (if torticollis is observed) are excellent sources for recovery. Sterile cotton-tipped swabs are preferred to wire inoculating loops, except for small organs in young birds, because swabs transfer more tissue. Several tissues should be cultured from each bird, as tissues with lesions do not always yield salmonellae. Portions of several tissues may be pooled and added to selective enrichment broth to increase the possibility of isolation (Fig. 2.1). Sections of the intestinal tract, especially the ceca and cecal tonsils, may be cultured by inoculation into selective enrichment broth. Serologic Reactors Because of the potential consequences involved in culturing serologic reactors, especially pullorum-typhoid, these cases should be cultured more intensively than routine clinical cases. Also, in some situations reactor birds may be asymptomatic, the salmonellae may be present in smaller numbers than in clinical cases, or they may be more localized. Recent use of antimicrobial agents may prevent the recovery salmonellae, although they may not completely clear the organism from the bird. Reactors should be evaluated by both direct and selective culture procedures (Fig. 2.1) (15,17). Tissues showing any pathologic lesions should be sampled using a sterile cotton-tipped swab and inoculated onto a nonselective plating medium. If desired the swab may be broken off into a tube of nonselective broth, incubated at 35-37 C for 24 hr, and plated onto nonselective agar. Portions of the liver, spleen, heart, gall bladder, ovaiy and oviduct should be pooled (10-15 g total) and then minced, blended, or stomached. Selective enrichment broth is added to the tissue (one part tissue homogenate to 10 parts broth) and incubated at 35-37 C for 24 hr, and if negative they are incubated for an additional 24 hr. After the internal organ samples have been removed, the intestinal samples may be taken. Portions of the ceca and the cecal tonsils (and other sections of the intestinal tract including the crop may added) should be pooled and minced, blended, or stomached. Selective enrichment broth is added to the sample (one part sample to 10 parts broth), incubated typically at 41 C for 24 hr, and plated on selective plating media.

CLINICAL DISEASE Most salmonellae are transmitted horizontally, but some serotypes (such as S. pullorum and S. enteritidis) can be transmitted vertically and often produce highly persistent flock infections. Pullorum disease primarily affects chickens, turkeys, and other fowls during the first few weeks of life. Fowl typhoid is also observed in mature poultry. These diseases are uncommon in pet birds species and pigeons. Paratyphoid salmonellae, particularly some strains of S. enteritidis, can occasionally cause morbidity or mortality in young birds of diverse species. Histopathologic lesions associated with avian salmonellosis include fibrinopurulent perihepatitis and pericarditis; purulent synovitis; focal fibmoid necrosis, lymphocytic infiltrations, and small granulomae in various visceral organs; and serositis of the pericardium, the pleuroperitoneum, and the serosa of the intestinal tract and mesentery (9,10,13). Infections with S. arizonae can produce encephalitis and hypopyon in young turkeys and chickens. SAMPLE COLLECTION

Both clinical tissue and environmental samples should be collected as aseptically as possible to prevent cross-contamination. This includes the use of sterile sampling materials (e.g., swabs, scoops, bags) and disposable gloves. It is critical that strict biosecurity 3

W. Douglas Waltman and Richard K. Gast

levels above 0.85 appear to promote the environmental survival and multiplication of salmonellae. The frequency and extent of sampling often varies according to the purpose of the monitoring. For example, the NPIP has established guidelines for testing breeding flocks for salmonellae (15). Environmental samples pose a challenge to the detection of the salmonellae, because salmonellae are often present in low numbers, salmonellae may be present but injured, and large populations of other bacteria are often present. Therefore, highly selective enrichment and plating media must be used. Drag Swabs. Flock infection or contamination can be conveniently detected by use of drag swabs. Properly assembled and sterilized 3or 4-square inch gauze pads moistened with DSSM are drawn by 34-ft lengths of cord across the surface of the floor litter or dropping pit for 15-20 min. The full length of the occupied building is traversed one or more times. The swabs are placed in individual sterile plastic bags at the farm and transported as soon as possible to the laboratory on ice packs. Selective enrichment broth is added directly to the bags containing the swabs (15). An alternative to the gauze pad is a commercially available sponge. Caution is advised to make sure these sponges are specially made for culturing purposes, because many household-type sponges contain bacteriocidal or bacteriostatic substances. In studies with broiler chickens, results from several unpooled drag swabs closely reflected intestinal excretion rates of salmonellae in the flock (7). Such sampling circumvents more cumbersome methods of environmental monitoring (5). Floor Litter. Samples should be collected from representative dry areas of floor litter; wet and caked ateas, including those around waterers and feeders, should be avoided because salmonellae survival may be poor in such areas. A sterile wooden tongue depressors or similar device is used to collect 5 g of dry litter from the upper 2.5-5.0 cm (1-2 inches) of floor litter into a sterile plastic bag from five sites in the pen or house. Additional sample pools should be collected from other areas of the house or pen to obtain the following ratio of samples to birds per pen: fewer than 500 birds, 5 pooled samples; 500-2500 birds, 10 pooled samples; 2500 birds or more, 15 pooled samples (15). The sample numbers suggested above are necessary for reasonable dependability in monitoring semimature or mature flocks in which excretion rates may be very low. Nest Boxes or Egg Belts. Loose litter in nests is a preferred sample site after breeder hens have been in lay 2-4 wk. Following the same procedure for the nest litter as given for the floor litter, collect fine material from the bottoms of at least five nests for each pooled sample. Samples need not be weighed or measured after some experience is acquired, but weight should not exceed 10-15 g per pooled sample. Many companies no longer use wood shavings in their nest boxes because of automated egg collection. In lieu of sampling nest shavings, swabbing the inside of an equal number of nest boxes using DSSM moistened gauze pads has been shown to be effective. Each pad may be used to sample 5-10 nest boxes and then placed into a sterile bag for culture. It is a good practice to combine sampling the floor, by using either litter or drag swabs, with nest sampling. In lieu of swabbing the nest boxes in houses with mechanical nests, the egg belts may be swabbed with DSSM-moistened swabs, usually at the front of the house. Dust. In some regions, dust has been a more dependable source for isolation of salmonellae than litter. Local experience should be a guide for determining the most dependable source. Samples are collected by DSSM moistened gauze pads or by wooden tongue depressors from 15 or more sites of dust accumulation per pen. Depending on the volume collected, five samples may be pooled into one. Use cotton-tipped swabs only for hard-to-reach areas.

One-Day-old Hatchlings It is often important to determine the presence of Salmonella contamination or infection in 1-day-old chicks or poults. Four methods have been used. The first method, approved by the National Poultry Plan(NPIP), takes 25 1-day-old chicks (in groups of 5) and pools the internal organs, yolk sac, and intestinal tracts from each group in three separate sterile plastic bags (16). Selective enrichment broth is added to all five groups of pooled samples. A second method cultures chick meconium collected during chick handling for sexing. Pools of about 5 g of meconium are collected from the chicks into sterile plastic bags and inoculated with selective enrichment broth. A third method cultures papers that line the boxes the chicks are transported in from the hatcheiy to the farm. The surface of these chick papers may be swabbed with double-strength skim milk (DSSM)-moistened gauze pads or by directly placing pieces of the paper into a sterile plastic bag and adding selective enrichment broth. A fourth method, with good sensitivity, uses 50-100 chicks that have been held for 48-72 hr with only clean water available for consumption. This promotes widespread dissemination of infection among boxmates and increases the likelihood of detection by culturing the paper liner or even by cloacal swabbing of the birds in some cases. However, an important difficulty in using this method is that the chicks must be maintained without cross contamination from external sources of Salmonella. Cloacal Swabs Cloacal swabs have been shown to be an unreliable means of detecting salmonellae from birds (4). Generally, salmonellae are excreted intermittently, and often in low numbers. Furthermore cloacal swabbing of birds requires laborious culture of 300-500 birds to provide dependable results. Egg Culture The surface of intact eggs can be sampled by immersion in 30-50 ml of selective enrichment broth in a plastic bag, with soaking or manual rubbing for at least 30 sec before removal of the egg. Eggshells can also be manually crushed in selective enrichment broth to allow more complete access of the media to internal surfaces. After disinfection of eggshells (generally in 70% ethanol or 2% tincture of iodine), egg contents can be cultured at a standard 1:10 ratio in selective or nonselective media. The contents of 10-30 eggs are often pooled for culturing because of the usual low incidence and level of Salmonella contamination. To allow salmonellae to multiply to easily detectable levels (and to minimize media consumption), pools of egg contents should be incubated for at least 1 day at 37 C or 3 days at 25 C before subculturing into broth media. Incubated egg pools can be transferred directly onto selective agar, but significantly greater detection sensitivity can be attained by using one or more enrichment steps. Environmental Monitoring Environmental monitoring or surveillance has become a useful method for predicting potential infection or colonization of flocks with the paratyphoid salmonellae. Environmental sampling has been shown to be effective and less invasive than other sampling methods. Although environmental monitoring may show good predictive evidence of salmonellae in a flock, it is an indirect indicator and flocks should not be diagnosed as infected based solely on the environmental sampling. The probability of detecting salmonellae in environmental samples depends on the number of salmonellae excreted into the environment by the flock, the survival of salmonellae in the sample locations, the intensity of sampling, and the culture methods used. Fecal shedding of salmonellae is often greatest during the first few weeks of life, when the susceptibility of chicks to infection is greatest. Litter surface water activity (equilibrium relative humidity) 4

Chapter 2

Cage Housing. The best methods of sampling for cage houses have not been fully determined. Methods in current use include sterile gauze pads moistened with DSSM to aseptically collect material from egg transport belts and elevators, rollers, diverters, and manure scrapers or cables. All segments of the house should be sampled. The drag swab method is also used with layer cages that allow droppings to accumulate under cages. Hatcheries. The area most likely to yield salmonellae is the hatcher during or following hatching. Selective Agar plates, opened and exposed for 5-10 min to the air in various parts of the hatcheiy building, have been used to monitor hatcheries. However, failure to find salmonellae on such plates is not a reliable indication of freedom from hatchery contamination. A sterile tongue depressor may be used to collect at least one heaping tablespoonful of fluff and dust from each hatcher. This sample is placed into a sterile plastic bag with selective enrichment broth. The most reliable sample has been the hatch residue (2). About 1015 g of eggshells and other hatch residue remaining after the chicks are removed from the hatch trays is placed into a sterile plastic bag to which selective enrichment broth is added. Alternatively, several hatch trays may be swabbed with gauze pads and pooled. Feed and Feed Ingredients. About 100 g of feed or feed ingredients should be collected into a sterile plastic bag representing multiple sites in a lot of feed. Care should be taken to prevent cross contamination of the samples. The Association of Official Analytical Chemists (AOAC) International method is the most widely accepted procedure for culturing feed and feed ingredients (1). Twenty-five grams of feed is mixed with 225 ml of lactose broth. The mixture is allowed to stand for 1 hr and the pH is adjusted to 6.8 + 0.2. The culture is incubated at 35 C for 24 hr. One milliliter of the culture is transferred into 10 ml of tetrathionate (TT) broth and 0.1 ml is transferred into Rappaport-Vassiliadis (RV) broth. The two enrichment broths are incubated at 43 C and 42 C, respectively. After 24 hr, the cultures are inoculated onto bismuth sulfite, Hektoen enteric (HE), and xylose lysine desoxycholate (XLD) agars. Some investigators have suggested that this procedure be modified by using universal preenrichment or buffered peptone water (BPW) as the preenrichment broth; by reducing the selective enrichment incubation temperatures to 41 C; and by replacing the plating media with more selective agars, such as brilliant green supplemented with novobiocin (BGN) and xylose lysine tergitol 4 (XLT4) agars.

Salmonellosis

PREFERRED CULTURE MEDIA AND SUBSTRATES

Recommended Steps, Media, and Incubation Times and Temperatures. The isolation and identification flow chart in Fig. 2.1 details the selection and use of preferred media in a variety of testing situations. Selective enrichment broths are typically inoculated at a 1:10 ratio of sample to broth, except for RV enrichment, which is inoculated at a 1:100 ratio from a pre-enrichment broth. Typically, internal organ samples and samples having lower background levels of bacteria may be incubated at 35-37 C. Intestinal and environmental samples having higher levels of background bacteria are commonly incubated at 40-43 C. Because of potential incubator problems and the sensitivity of some strains to higher temperatures, the preferred incubation temperature is 41+0.5 C. Routine monitoring of the temperature of incubators is critical when using the higher temperatures. Selective enrichment broths should be incubated 20-24 hr and plated. Use of a delayed secondary enrichment (DSE) procedure is highly recommended (see below), but if not, the enrichment broths should at least be incubated an additional 24 hr and replated. Precautions for Environmental and Intestinal Samples Isolation of salmonellae from environmental and intestinal monitoring samples is much more demanding than isolation from samples collected from bacteremic (clinically affected) or carrier (serologic reactor) birds. In bacteremic or carrier birds, salmonellae populations are often high, particularly with respect to bacterial competitors. In contrast, salmonellae levels are ordinarily low in environmental or intestinal samples where competing bacteria are found in high levels. These competitors, if not controlled, seriously impair media detection efficiency, resulting in false negatives. The dependable isolation of salmonellae from environmental and intestinal samples is, therefore, significantly improved by incubation of enrichment broths at 41 + 0.5 C for a full 24-36-hr period, use of novobiocin- or tergitol (niaproof) 4-supplemented media, DSE of primary selective enrichment broths, and use of a plate-streaking technique that produces well-separated colonies.

Non-Selective Media Salmonellae grow well on such nonselective broth media as veal infusion, brain-heart infusion, or nutrient broth. Nonselective agars that may be used include blood agar and nutrient agar. MacConkey agar, although mildly selective, is frequently used as a nonselective medium. These media are preferred for tissues from clinical cases and serologic reactors in which the likelihood of competing bacteria is low. Use of nonselective media is important when S. pullorum or S. gallinarum are suspected, as some strains may be inhibited by selective media.

Shipping and Storing Samples Holding Media. Studies have shown that the best moistening agent and holding media for environmental swabs is DSSM (12). This medium is prepared by dissolving 200 g Bacto Skim Milk (BD Diagnostics Systems, Sparks, MD) in 1 liter distilled or deionized water in a large flask and autoclaving. The DSSM may be stored in the refrigerator. Sample Storage Times and Temperatures. Specimens from clinical diagnostic consignments and serologic reactors should be cultured immediately upon collection and no more that 24 hr after collection even if they are refrigerated. Meconium, freshly voided feces, and water should be refrigerated at Ί-b C as soon as possible and inoculated into appropriate culture media within 24 hr. Drag swabs and other swabs containing DSSM may be refrigerated for 3 days or frozen for 7 days before inoculation of selective enrichment broth (12). Because the survival rate of salmonellae in floor litter samples varies considerably, even under refrigeration, it is best to culture floor litter within 2 days of collection. This is advisable despite the fact that some dry environmental samples have yielded salmonellae when held for several months at room temperature and low humidity.

Pre-enrichment Preenrichment is the process of inoculating nonselective broth media with the sample and incubating it for 24 hr at 35-37 C. Typically, 1.0 ml of the preenriched culture is transferred to 10 ml of a TT enrichment broth or 0.1 ml is transferred to 10 ml of RV enrichment broth. The purpose of preenrichment is to revive injured salmonellae that may be present in some samples. Typical preenrichment media include lactose broth and BPW. Selective Enrichment Broths Tetrathionate Enrichments. Tetrathionate broths are the preferred selective enrichments for salmonellae isolation. Older formulations, such as Mueller-Kauffrnann TT brilliant green broth, require the addition of both iodine and brilliant green solutions to the broth immediately before use. However, newer formulations, such as TT broth, Hajna, or TT, Hajna and Damon (BD Diagnostic Systems,

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Sparks, MD) are supplied already containing brilliant green and require only the addition of an iodine solution to the broth base. The newer TT formulations contain more nutrients and buffers. The iodine solution differs for the individual TT formulations, so care must be taken to ensure the appropriate one is used and that it is prepared and stored correctly. Selenite Enrichments. The use of selenite enrichments are no longer recommended because they have a short shelf life, are not as effective at higher incubation temperatures (3), potentially mutagenic (11), and in many geographic areas are considered to be a hazardous waste. Rappaport-Vassiliadis Enrichments. A selective enrichment that is gaining acceptance and appears to be replacing selenite enrichments is RV broth. The RV enrichment procedure begins with preenrichment of the sample in BPW and then inoculation into RV broth at a ratio of 1:100. The RV broth should be incubated at 4142 C.

Xylose Lysine Desoxycholate, XLD Supplemented with Novobiocin, and XLT4 Agars. The differential characteristics of these media include the lysine decarboxylation and H2S-producing abilities of most salmonellae. Hydrogen sulfide-positive salmonellae colonies are nearly or fully jet black. Proteus (inhibited on XLDN and XLT4) colonies may appear blackish, but the color is less brilliant, tending to have a gray or green cast as the colonies age. Colonies of Citrobacter freundii ordinarily have black centers and prominent creamy-colored borders on these media. Other Plating Media. Bismuth sulfite and Hektoen enteric agars, although widely used in human clinical situations and advocated by the Food and Drug Administration (FDA) and AOAC International for food and feed, lack the sensitivity and specificity of BGN and XLT4. Modified lysine iron agar (MLIA) incorporates novobiocin and is similar in recovery to BGN or XLDN. Rambach agar, a new chromogenic agar, appears to be effective, but it is more expensive than other options. Recently, several other chromogenic agars have been developed for the isolation of Salmonella. The most thoroughly studied of these, Miller-Mallinson (MM) media, has been found to be very effective.

Delayed Secondary Enrichment (DSE) After the 24-hr incubation of the TT and plating onto BGN and XLT4 agars, the TT culture is left at room temperature for 5-7 days. If the original plating was negative for salmonellae, 0.5-1.0 ml of the original TT broth is transferred into 10 ml of fresh TT broth and incubated at 37 C for 24 hr. The new TT broth is plated and processed as before (16). Studies have shown that DSE increases the recovery rates of salmonellae from most sample types (18).

Rapid Salmonella Detection Techniques A number of rapid tests have been approved for detecting salmonellae in various samples. Typically these have been approved based on comparison to AOAC International culture procedures (1). The rapid tests have several advantages over conventional culture, such as obtaining results in less than 48 hr, being less labor intensive, and having the possibility of automation. For many sample types, especially clinical, processing plant, and food samples, these rapid tests appear to be veiy effective. However, some sensitivity and specificity problems are apparent with feed and environmental samples. A variety of rapid detection systems are commercially available, including antigen-capture enzyme-linked immunosorbent assay (AC-ELISA) systems, DNA probes, polymerase chain reaction systems, immunodiffusion, immunofluorescence, magnetic bead systems, bioluminescence, and other novel applications. A listing of these tests may be found at the FDA-BAM website. Currently, NPIP has approved a rapid ruthenium-labeled sandwich immunoassay and two PCR systems for detecting Salmonella in environmental samples (15). As technology advances the sensitivity and specificity of rapid systems will surely increase and allow more widespread use.

Selective Plating Media Rationale for Using. Because of the high levels of nonsalmonellae bacteria in most intestinal and environmental samples, selective plating media must be used to assist the selective enrichment broth in inhibiting other bacteria. Conventional salmonellae plating media lacks the selectivity necessary for these samples. Many media can inhibit most coliforms, however, the primary problems are with species of Proteus, Providencia, Morganella, and Pseudomonas. These bacteria resemble salmonellae on some plating media and must be screened, resulting in high percentages of false-positive cultures. The addition of 20 pg of novobiocin (N1628 Sigma Chemical Co., St. Louis, Mo.) per ml of plating media, especially brilliant green (BG) (14) and XLD agars, results in the inhibition of Proteus. Another plating medium that has proven to be effective is XLT4 (BD Diagnostic Systems, Sparks, MD). This medium inhibits Proteus and Pseudomonas aeruginosa. A combination of two plating media that are predicated on different selective and differential characteristics is advocated. BGN and XLT4 are two good choices (8). The use of BGN agar, which demonstrates the positive hydrogen sulfide (H2S) production characteristic, increases the likelihood of detecting atypical strains. Hydrogen sulfide­ negative strains undetected on XLT4 should be obvious on BGN, because they usually remain lactose-negative. Conversely, lactose­ positive strains (eg., S. arizonae and others) undetected on BGN should be obvious on XLT4 because they usually remain MS­ positive. Brilliant Green (BG) Supplemented with Sulfapyridine or Sulfadiazine (BGS), and BGN Agars. Brilliant green agar has been supplemented with antimicrobial compounds to make it more selective. The purpose of adding sulfapyridine or sulfadiazine and novobiocin was primarily to inhibit Proteus species. Salmonellae colonies on these media are usually transparent pink to deep fuchsia, surrounded by a reddish medium. These colonies may lose this characteristic appearance if there is a heavy growth of lactosefermenting colonies or if the colonies are not well separated. Some nonsalmonellae bacteria produce colonies similar in appearance to salmonellae and must be screened further. Host-adapted S. pullorum and 5. gallinarum colonies are smaller and grow slower than nonhost-adapted salmonellae. Consequently, all plates should be incubated for 48 hr.

AGENT IDENTIFICATION Basic Identification Screening Media The combined use of TSI and LI agar slants is generally sufficient for presumptive identification of most salmonellae-suspect colonies. At least three well-separated colonies are selected for transfer to each set of TSI and LI agar slants. Each colony is stabbed into the butt of the TSI and LI agars and streaked across the slants. The tubes are read after incubation for 24 hr at 37 C. The use of more selective plating media (BGN and XLT4) to screen these salmonellae suspect colonies results in fewer false positives (nonsalmonellae bacteria). The presence of multiple serotypes may be more likely in some samples than in others. Also, in situations where S. enteritidis, S. pullorum, qt S. typhimurium are being monitored, and other salmonellae are present, screening many more colonies may be necessary to ensure the absence of S. enteritidis, S. pullorum, or S. typhimurium serotypes. The Colony Lift Immunoassay (Synbiotics Corp., San Diego, CA) may be useful in detecting group D salmonellae in these situations (6). Triple Sugar Iron (TSI) Agar. Most salmonellae produce an alkaline (red) slant and acid (yellow) butt, with gas bubbles in the agar and a blackening due to H2S production that often obscures the

6

Chapter 2

acid reaction in the butt of the tube. Salmonella gallinarum does not form gas in TSI, whereas S. pullorum may show weak gas production. Both of these Salmonella may or may not show H2S production. Some paratyphoid strains may be H2S-negative in this medium. Lysine Iron (LI) Agar. Salmonellae will show lysine decarboxylation, with a deeper purple (alkaline) slant and alkaline or neutral butt with slight blackening due to H2S production, with the exceptions noted in the previous paragraph. LI agar is useful in differentiating common intestinal flora such as Proteus and Citrobacter from further consideration. Proteus (also Providencia and Morganella) produces a reddish or port-wine colored slant, indicating lysine deamination, and a yellow (acid) butt on LI agar medium. Citrobacter gives a purple (alkaline) slant and a yellow (acid) butt, with some H2S production. Rarely, both slant and butt are yellow.

Salmonellosis

SEROLOGIC DETECTION IN THE HOST

Serologic Testing of Poultry Flocks Several serologic tests have been developed for detecting antibodies to salmonellae. The most commonly used tests for pullorum-typhoid include the whole blood plate (WBP) (not approved for use in turkeys), rapid serum plate, standard macroscopic tube agglutination, microagglutination, and microantiglobulin tests (9,15). Possibly the most widely used of these tests is the WBP test, which can be performed in the field using commercially available antigens. A drop of fresh blood is mixed with a drop of antigen on a plate, mixed, and observed for agglutination within 2 min. The tube agglutination test is also commonly used. It requires sera for testing with an antigen that may be obtained from the NVSL. In the macroscopic tube agglutination test the sera are diluted 1:25 to 1:50 for chickens or 1:25 for turkeys. The sera are mixed with the antigen, incubated at 37 C for 20-24 hr, and observed for agglutination. Because the WBP test is not approved for turkeys, the tube agglutination or rapid serum agglutination tests are typically used. Agglutination tests have also been used for detecting antibodies to various paratyphoid salmonellae, especially S. typhimurium, S. arizonae, and S. enteritidis. These tests have met with varying degrees of success due to sensitivity and specificity problems and thus have not found widespread commercial application. Because S. enteritidis and S. pullorum share somatic antigens, pullorumtyphoid antigen preparations have sometimes been applied for detecting antibodies to S. enteritidis. Enzyme-linked immunosorbent assays (ELISAs) have been developed for detecting antibodies to various salmonellae. The most widely used ELISAs have been developed for detecting antibodies to S. enteritidis. These ELISAs have been used with some success, particularly in Europe, but concerns about their specificity still persist. The various serologic tests for detecting antibodies to salmonellae, especially the agglutination tests, are subject to false positive reactions. Confirmation of all positive serologic tests by culturing the bird for salmonellae is important.

Biochemical and Serological Confirmation Biochemical Identification. Further biochemical tests may be necessary to confirm an isolate as Salmonella. Table 2.1 lists several typical biochemical reactions of salmonellae. Key tests for differentiating salmonellae from other bacteria include those for urea {Proteus and most Providencia and Morganella are urease positive), beta-galactocidase (salmonellae with the exception of S. arizonae, are negative whereas C. freundii and other coliforms are positive), and indole (Escherichia coli and Escherichia tarda produce indole, salmonellae do not). Commercially available identification systems may also be very useful in confirming isolates. They use a number of biochemical tests in a simple to use format. Serological Typing. The presumptive salmonellae colonies identified by the TSI and LI agar reactions should be serologically typed. The first phase of serologic typing is to serogroup the isolates based on their somatic O-group antigens using commercially available polyvalent somatic antisera. Individual polyvalent antisera are tested against each isolate in a simple plate agglutination assay. Bacterial growth from an agar plate is emulsified in a small amount of physiologic saline to form a milky suspension, and one drop of polyvalent O antisera is mixed with it on a slide or plate. The agglutination reaction is read within 60 sec. After determining which polyvalent antiserum agglutinates with the isolate, additional testing is performed using individual single factor O-group antisera that comprised the polyvalent antiserum. In this maimer, each isolate is typed to a specific serogroup. The next step, which is often performed at a reference laboratory, is to determine the serotype. The serotype is based on the flagellar antigen present on almost all salmonellae, except 5. pullorum and S. gallinarum. Commercially produced antisera are available for serotyping most isolates (BD Diagnostic Systems, Sparks, MD). The serotyping procedure involves an antigen extraction step with formalin, followed by a microtube agglutination test.

DIFFERENTIATION FROM CLOSELY RELATED AGENTS Young birds with generalized Salmonella infections may show signs and lesions identical to any bacteremia. In severe outbreaks of salmonellosis, fibrinous liver and heart lesions may be very similar to those seen with air sac disease (colibacillosis). The heavy yellowish white cheesy exudate covering the retina of turkey poults with S. arizonae infection and occasionally with other paratyphoid infections can be confused with signs of aspergillosis. Nervous signs associated with infection of the brain of fowl may resemble those of Newcastle disease or other diseases affecting the central nervous system. Joint involvement may be mistaken for synovitis or bursitis due to other infectious agents.

7

W. Douglas Waltman and Richard K. Gast

Figure 2.1 General Salmonella /solation and identification

A Hajna TT or Mueller-Kauffmann tetrathionate enrichment broths. For RV broth, follow special inoculation and preenrichment instructions of manufacturer B Beef extract, veal infusion, or comparable non-selective media. A broth detect can help detect low Salmonella levels in live birds. C BGN in combination with XLT4 is preferred (refer to text) D Colony lift immunoassays can significantly increase the reliability detecting Group D Salmonella (S. enteritidis, S. pullorum, etc. on plating agars (refer to text) E If combined results with TSI and LI agars, additional identification media, and O-group screening procedures are inconclusive, restreak original colony onto selective plating agar to check for purity. F Reevaluate if epidemiologic, necropsy or other information strongly suggests the presence of an unusual strain of Salmonella.

8

Chapter 2 Table 2.1 Typical biochemical reactions of Salmonella K Paratyphoid S. pullorum S. gallinarum salmonellae Dextrose A A(G)b AG Lactose Sucrose Mannitol A A(G) AG Maltose A AGe (-)D Dulcitol A AG Malonate Urea + Motility -

Media

Salmonellosis

4. Fanelli, M J., W. W. Sadler, C. E. Franti, and J. R. Brownell. Localization of salmonellae within the intestinal tract of chickens. Avian Dis. 15:366-375. 1971. 5. Kingston, D. J. A comparison of culturing drag swabs and litter for identification of infections with Salmonella spp. in commercial chicken flocks. Avian Dis. 25:513-516. 1981. 6. Lamichhane, C. M, S. W. Joseph, W. D. Waltman, T. Secott, E. M Odor, J. deGraft-Hanson, E. T. Mallinson, V. Vo, and M Blankford. Rapid detection of Salmonella in poultry using the colony lift assay. In: Proceedings of the Southern Poultry Science Society. Atlanta, Ga. Poult. Sci. (Suppl. 1)74:198. 1995. 7. Mallinson, E. T., C. R. Tate, R. G. Miller, B. Bennett, and E. RussekCohen. Monitoring poultry farms for Salmonella by drag swabs sampling and antigen capture immunoassay. Avian Dis. 33:684—690. 1989. 8. Miller, R. G., C. R. Tate, E. T. Mallinson, and J. A. Scherrer. Xylose lysine tergitol 4: an improved selective agar medium for the isolation of Salmonella. Poult. Sci. 70:2429-2432. 1991. 9. Gast, R. K. Paratyphoid infections. In: Diseases of poultry, 11th ed. Y. M Saif, H. J. Barnes, J. R. Glisson, A M Fadly, L. R. McDougald, and D. E. Swayne, eds. Iowa State University Press, Ames, Iowa. pp. 583-613. 2003. 10. Nagaraja, K. V., B. S. Pomeroy, and J. E. Williams. Arizonosis. In: Diseases of poultry, 9th ed. B. W. Calnek, H. J. Bames, C. W. Beard, W. M Reid, and H. W. Yoder, Jr., eds. Iowa State University Press, Ames, Iowa, pp. 130-137. 1991. 11. Noda, Μ, T. Takano, and H. Sakurai. Mutagenic activity of selenium compounds. Mutat. Res. 66:175-179. 1979. 12. Opara, Ο. O., L. E. Carr, C. R. Tate, R. G. Miller, E. T. Mallinson, L. E. Stewart, and S. W. Joseph. Evaluation of possible alternatives to double­ strength skim milk used to saturate drag swabs for Salmonella detection. Avian Dis. 38:293-296. 1994. 13. Shivaprasad, Η. N. Pullorum Disease and Fowl Typhoid. In: Diseases of poultry, 11th ed. Y.M Saif, H. J. Bames, J; R. Glisson, A. M Fadly, L. R. McDougald, and D. E. Swayne, eds. Iowa State University Press, Ames, Iowa. pp. 568-582. 2003. 14. Tate, C. R., R. G. Miller, E. T. Mallinson, and L. W. Douglass. The isolation of salmonellae from poultry environmental samples by several enrichment procedures using plating media with and without novobiocin. Poultry Sci. 69:721-726. 1990. 15. United States Department of Agriculture (USDA). National Poultry Improvement Plan and Auxiliary Provisions. Animal and Plant Health Inspection Service, 91-55-063. USDA, Washington, D.C. Feb 2004. 16. Waltman, W. D., A. M Home, C. Pirkle, and T. G. Dickson. Use of delayed secondary enrichment for the isolation of Salmonella in poultry and poultry environments. Avian Dis. 35:88-92. 1991. 17. Waltman, W. D., and A. M Home. Isolation of Salmonella from chickens reacting in the pullorum-typhoid agglutination test. Avian Dis. 37:805-810. 1993. 18. Waltman, W. D., A. M Home, and C. Pirkle. Influence of enrichment incubation time on the isolation of Salmonella. Avian Dis. 37:884—887. 1993.

S. arizonae

AG (-)C AG AG +F +

A = acid produced; G = gas produced; 0 = variable reaction B Blood agar is generally used as the non-selective agar plate. If desired, the swabs or pieces of tissue may be inoculated into a non-selective broth, such cas brain-heart infusion or nutrient broth. Brilliant green agar supplemented with novobiocin (BGN) in combination with xylose-lysine-tergitol (niaproof) 4 (XLT4) agar are preferred (refer to text). If S. pullorum is suspected, MacConkey (MAC) agar may be substituted for XLT4 agar). D Colony lift immunoassays may increase the reliability of detecting Group D Salmonella (e.g. S. enteritidis, S. pullorum, etc) on plating media (refer to text). E Salmonella typhimurium var. Copenhagen, as isolated from pigeons, frequently forms no acid in maltose broth and can be confused with S. pullorum. However, it is motile. F Turns deep Prussian blue within 24 hr.

ACKNOWLEDGMENT

The guidance supplied by Edward T. Mallinson is very gratefully acknowledged REFERENCES

1. Association of Official Analytical Chemists (AOAC) International. Official methods of analysis, 16th ed. P. Cunniff, ed. AOAC International, Gaithersburg, Md. pp. 55-94. 1996. 2. Bailey, J. S., N. A. Cox, and Μ E. Berrang. Hatchery-acquired salmonellae in broiler chicks. Poult. Sci. 73:1153-1157. 1994. 3. Carlson, V. L., and G. H. Snoeyenbos. Comparative efficacies of selenite and tetrathionate enrichment broths for the isolation of Salmonella serotypes. Am. J. Vet. Res. 35:711-718. 1974.

9

3 COLIBACILLOSIS Margie D. Lee and and Lisa K. Nolan SUMMARY. Escherichia coli is the causative agent of colibacillosis in poultry. The disease results from a systemic infection involving the blood, joints, and/or air sacs of birds. Young birds (4-8 wk old) may die of acute septicemia that is preceded by only a brief period of anorexia and depression. At necropsy, lesions are sparse but may include swollen, dark-colored liver and spleen and increased fluid in the body cavities. Birds surviving the acute phase may develop fibrinopurulent airsacculitis, pericarditis or arthritis. Whether lesion-associated isolates are primary pathogens or whether environmental factors are responsible is unknown at this time. Agent Identification. Isolation of E. coli is significant if made from the internal organs or blood from fresh carcasses. MacConkey agar is selective and differential for E. coli and is preferred for primary isolation. Presumptive positive colonies (pink on MacConkey that produce A/A reaction on triple sugar iron agar and that are oxidase-negative, gram-negative rods should be confirmed as E. coli by a positive indole test and the inability to produce H2S. But some isolates do not form pink colonies on MacConkey, therefore multiple colonies should be separately inoculated on triple sugar iron agar to detect these pathogens. Serologic Detection in the Host. Host serology is not useful for diagnosis because many birds have antibody to normal intestinal flora isolates of E. coli.

fibrinopurulent lesions suggest subacute colibacillosis, swab samples of exudate should be collected from the pericardial sac, air sacs, and joints. Lesions present more than 1 wk are often sterile. When postmortem changes are obvious, bone marrow samples may be useful because they are less likely than other tissues to contain intestinal E. coli. Escherichia coli isolates survive well on sealed agar slants for storage and shipping. For long-term storage, mix E. coli broth culture with sterile glycerol 1:1 and store at -20 to -60 C.

INTRODUCTION Colibacillosis of poultry is a common systemic infection caused by avian pathogenic Escherichia coli (APEC). The disease is economically important to poultry production worldwide. Colibacillosis occurs in many forms. It may occur as colisepticemia, which typically leads to death. However, some birds fully recover from colisepticemia, and others may recover with sequelae (2). Colibacillosis may also be localized, manifesting as omphalitis, yolk sac infection, cellulitis, swollen head syndrome, enteritis, acute vaginitis, salpingitis, or peritonitis (2,12). For E. coli infections to become clinically apparent, adverse environmental factors or other infectious agents are usually required (2, 3). A foodbome link between human disease and APEC has not been established. However, recent reports of similarities between the virulence attributes of APEC and human extraintestinal pathogenic E. coli, suggest that some APEC may be capable of causing disease in human beings (4,10).

PREFERRED CULTURE MEDIA AND SUBSTRATES

Escherichia coli grows well in most commonly used culture media, but differential media are useful for primary isolation. MacConkey's agar is a selective and differential medium for the isolation of enteric organisms. Tryptose blood agar with 5% bovine blood can be used as a primary culture medium to support growth of E. coli and other bacterial pathogens. Several differential biochemical characteristics can be obtained through use of Kligler’s iron agar or triple sugar iron agar slants. All the above media are readily available from BD Diagnostics (Sparks, Md.). Microbiology identification kits such as the API 20E (bioMerieux Vitek, Hazelwood, Mo.) and Enterotube (Becton Dickinson Microbiology Systems) are useful for performing the biochemical tests and are available commercially.

CLINICAL DISEASE Clinical signs of colibacillosis are nonspecific and vaiy with the age of bird, duration of infection, organs involved, and concurrent disease conditions. In young (4 to 8-wk-old) broilers and poults dying of acute septicemia, death is preceded by a brief period of anorexia, inactivity, and somnolence. At necropsy, lesions are sparse except for swollen, dark-colored liver and spleen and increased fluid in all body cavities. Birds surviving the septicemia phase of the disease are unthrifty and develop subacute fibrinopurulent airsacculitis, pericarditis, perihepatitis, and lymphocyte depletion in the bursa and thymus. Airsacculitis,a classic lesion of colibacillosis, occurs following respiratory exposure to large numbers of E. coli, but also occurs as a sequel to bacteremia. Other less common lesions are arthritis, osteomyelitis, salpingitis, and pneumonia (1,3,6). Cellulitis, responsible for substantial economic losses for the poultry industry, has also been associated with E. coli infection (12).

AGENT IDENTIFICATION

Colony Morphology and Biochemical Features Blood samples should be diluted 1:10 in brain-heart infusion broth then inoculated directly onto MacConkey’s plates. Swabs from lesions can be used to streak MacConkey’s plates. MacConkey’s agar should be incubated aerobically for 18-24 hr at 37 C for the primary isolation of E. coli. Escherichia coli, Enterobacter, and Klebsiella ferment lactose and can be distinguished from the other enteric organisms. On MacConkey's agar, most E. coli and some Enterobacter isolates produce 1-2-mm-diameter hot-pink colonies (lactose positive) whereas Klebsiella and Enterobacter aerogenes form large mucoid pink colonies (8). However slow lactosefermenting E. coli are frequently isolated from cases of colibacillosis and these may not form pink colonies. If primary cultures reveal large numbers of a predominant colony type suggestive of E. coli, pick several of these colonies and use them to separately inoculate triple sugar iron agar (or Kligler’s iron agar), sulfide-indole-motility (SIM) medium, and blood agar plates. If the clinical signs suggest colibacillosis infection, and no hot-pink colonies are seen on the MacConkey’s plates, pick a couple of non­ pink colonies to inoculate blood agar, triple sugar iron agar and SIM. A Gram stain and the oxidase reaction can be performed on colonies from blood agar plates. Escherichia coli, Enterobacter,

SAMPLE COLLECTION Only internal organs or blood, not feces or intestine, are useful samples. Because normal intestinal flora E. coli readily invade other tissues after death, specimens from fresh carcasses are necessary. When acute colisepticemia is suspected, heart blood and liver should be sampled aseptically. One ml of blood collected by needle and syringe can be used to inoculate broth media (1:10), which is used to streak agar plates. Sterile culture swabs or inoculation loops can be stabbed into the liver parenchyma after searing the capsule with a flamed scalpel or spatula. When

10

Chapter 3 Colibacillosis

and Klebsiella are oxidase-negative, gram-negative rods. On triple sugar iron agar (or Kligler’s iron agar), these organisms produce acid (even the slow lactose-fermenting E. coli) and gas but not H2S. On SIM medium, E. coli is positive for the indole reaction, positive or negative for motility, and negative for H2S, whereas Enterobacter and Klebsiella are usually indole-negative (Fig. 3.1). The site of sample collection, condition of the carcass, and nature of the lesion are important when deciding whether isolation of E. coli is relevant. The isolation of a pure culture of E. coli from the organs or blood of moribund birds or freshly dead carcasses is indicative of colibacillosis. Pathogenicity of E. coli isolates has traditionally been established by inoculating young (less than 3-wkold) chicks or poults parenterally with 0.1 ml of overnight broth culture. Pathogenic isolates should produce death or characteristic lesions of colibacillosis within 3 days. Alternately, inoculation of embryonated eggs may be used to establish the pathogenicity of E. coli isolates (7). Also, many APEC share a complex of plasmidassociated virulence genes, whose presence may be helpful in distinguishing APEC from non-pathogenic strains (9). Hence, multiplex PCR, targeting these common genes, may have value in confirming the pathogenic nature of E. coli isolates (11).

Antimicrobial Susceptibility APEC isolates are frequently resistant to more than one antibiotic. Seventy to 90% of isolates are resistant to sulfa drugs, tetracyclines, streptomycin, and gentamicin. It is not uncommon to find isolates that are multi-resistant to greater than 3 antibiotics. Resistance to fluoroquinolones is less frequently detected; a recent study (13) reported that 84% of isolates were susceptible to enrofloxacin.

SEROLOGIC DETECTION IN THE HOST Serology is not commonly used to detect E. coli infection.

DIFFERENTIATION FROM CLOSELY RELATED AGENTS Colibacillosis should be distinguished from other bacterial infections causing fatal septicemia or fibrinopurulent inflammation of air sacs, pericardium, joints, and other viscera. Diseases to be considered in the differential diagnosis include mycoplasmosis, salmonellosis, pasteurellosis, pseudotuberculosis, erysipelas, chlamydiosis, and staphylococcosis. Colibacillosis is a common complication of concurrent viral respiratory or enteric infections. REFERENCES

1. Arp, L.H.

Pathology of spleen and liver in turkeys inoculated with

Escherichia coli. Avian Pathol. 11:263-279. 1982.

2. Barnes, H.J., J.-P. Vaillancourt, and W.B. Gross. Colibacillosis. In: Diseases of poultry, 11th ed Y.M Saif, H. J. Bames, J.R. Glisson, A.M Fadly, L.R. McDougald, and D.E. Swayne, eds. Iowa State University Press, Ames, Iowa. pp. 631-652. 2003. 3. Gross, W.B. Colibacillosis. In: Diseases of poultry, 9th ed. B.W. Calnek, H.J. Bames, C.W. Beard, W.M Reid, and H.W. Yoder, eds. Iowa State University Press, Ames, Iowa. pp. 138-144. 1991. 4. Johnson, J.R., A.C. Murray, A. Gajewski, M Sullivan, P. Snippes, MA. Kuskowski, and K.E. Smith. Isolation and Molecular Characterization of nalidixic acid-resistant extraintestinal pathogenic Escherichia coli from retail chicken products. Antimicrob Agents Chemother. 47: 2161-2168. 2003. 5. Kaper, J. B., J.P. Nataro, and H.L.T. Mobley. Pathogenic Escherichia coli. Nature Rev. Microbiol. 2: 123-140. 2004. 6. Nakamura, K., M Maecla, Y. Imada, T. Imada, and K. Sato. Pathology of spontaneous colibacillosis in a broiler flock. Vet. Pathol. 22:592-597. 1985. 7. Nolan, L.K., R.E. Wooley, J. Brown, K.R. Spears, H.W. Dickerson, and M Dekich. Comparison of a complement resistance test, a chicken embryo lethality test, and the chicken lethality assay for determining virulence of avian Escherichia coli. Avian Dis. 36:395-397. 1992. 8. Quinn, P.J., ME. Carter, B.K. Markey, and G.R Carter. Clinical veterinary microbiology. Mosby-Year Book Limited, London, England, pp. 209-236. 1994. 9. Rodriguez-Siek, K.E., C.W. Giddings, M Fakhr, C. Doetkott, T.J. Johnson, and L.K. Nolan. Characterizing an APEC pathotype. Vet Res. 36: 1-16. 2005. 10. Rodriguez-Siek, K.E., C.W. Giddings, T.J. Johnson, M Fakhr, C. Doetkott, and L.K. Nolan. Comparison of Escherichia coli implicated in human urinary tract infection and avian colibacillosis. Microbiol. 151: 2097-2110. 2005. 11. Skyberg, J.A, S.M Home, C.W. Giddings, R.E. Wooley, P.S. Gibbs, and L.K. Nolan. Characterizing avian Escherichia coli isolates with multiplex PCR Avian Dis. 47:1441-1447,2003. 12. Vaillancourt, J.-P., and H.J. Bames. Coliform Cellulitis. In: Diseases of poultry, 11th ed. Y.M Saif, H. J. Bames, J.R Glisson, A.M Fadly, L.R. McDougald, and D.E. Swayne, eds. Iowa State University Press, Ames, Iowa. pp. 652-656. 2003. 13. Zhao, S., J.J. Maurer, S. Hubert, J.F. De Villena, P.F. McDermott, J. Meng, S. Ayers, L. English, and D.G. White. Antimicrobial susceptibility and molecular characterization of avian pathogenic Escherichia coli isolates. Vet. Microbiol. 107:215-24,2005.

Figure 3.1. Escherichia coli identification scheme .Diagnosis of colibacillosis is dependent on distinguishing pathogenic isolates from nonpathogenic, normal intestinal flora isolates of E. coli.

Serogrouping of Isolates Escherichia coli serogroups are based on O antigens; but serotypes of E. coli are based on O antigens, as well as flagellar and/or capsular antigens (5). There is great diversity of serogroups/types among APEC, although some occur more commonly than others (e.g., 078 and 02), and many isolates are not typable (9). Routine serotyping of APEC is usually impractical, but for epidemiologic studies, isolates can be serotyped for a fee by the Gastroenteric Disease Center (Pennsylvania State University, University Park, Penn.).

11

4 PASTEURELLOSIS, AVIBACTERIOSIS, GALLIBACTERIOSIS, RIEMERELLOSIS, AND PSEUDOTUBERCULOSIS John R. Glisson, Tirath S. Sandhu, and Charles L. Hofacre

SUMMARY. In avian hosts, certain members of the genera Pasteurella, Avibacterium, Gallibacterium and the species Riemerella anatipestifer and Yersenia pseudotuberculosis cause septicemic and respiratory disease. Acute diseases caused by these Gram-negative bacteria are often characterized by high flock mortality and morbidity. Clinical signs of acute disease include ruffled feathers, depression, increased respiratory rate, cyanosis, emaciation, and diarrhea. Lesions may consist of hemorrhages, swollen liver and spleen, focal necrotic areas in the liver, airsacculitis, and increased pericardial and peritoneal fluids. With R. anatipestifer infection, common gross lesions in ducks are fibrinous pericarditis, hepatitis and airsacculitis. Chronic disease signs produced by Pasteurella sp., R. anatipestifer and Y. pseudotuberculosis can include ocular and nasal discharge, swelling of tissues such as joints and wattles, and stunted growth. Pasteurella This genus includes P. multocida, the cause of fowl cholera. Two similar organisms, Avibacterium gallinarum (formerly Pasteurella gallinarum) and Gallibacterium anatis biovar haemolytica (formerly Pasteurella haemolytica) are include in this chapter because they can be isolated from the respiratory tract and have been associated with respiratory disease in poultry (1,4). Five serogroups of P. multocida (A, B, D, E and F) have been isolated from avian species, but serogroup A strains are the major cause of fowl cholera. Sixteen somatic serotypes may occur among strains of P. multocida. However, somatic serotypes 1, 3 and 4 are most commonly isolated. Agent Identification. Pasteurellas are identified by cell and colony morphology, Gram stain, and reactions in biochemical and other tests. A mucopolysaccharidase test can be used for presumptive identification of P. multocida serogroups A, D and F. Gel diffusion precipitin tests are used to determine somatic serotype. Serologic Detection in the Host. Serologic tests are not commonly used to detect infections by Pasteurella sp. in poultry.

Riemerella anatipestifer Riemerella anatipestifer infection occurs in ducklings usually at 1-8 wk of age. Primary isolations of R. anatipestifer are made on blood or trypticase soy agar incubated at 37C in a candle jar or CO2 incubator. At least twenty one serotypes have been identified using agglutination tests. Agent Identification. Riemerella anatipestifer is identified by cell and colony morphology, Gram stain, and biochemical characteristics. Fluorescent-antibody technique can be used to detect and identify R. anatipestifer in tissues or exudates from infected birds. Serologic Detection in the Host. Serum antibodies against R. anatipestifer can be detected in poultry by ELISA. Yersenia pseudotuberculosis Pseudotuberculosis, caused by infection with Y pseudo tuberculosis, is infrequently reported in poultry. Selective media are used for isolation of the bacterium from feces. There are six serotypes (I-VI) based upon heat-stable antigens using agglutination and agglutination­ adsorption tests. Among strains isolated from birds, serotype I is the most common. Serotypes V and VI have not been reported in birds. Agent Identification. Yersenia pseudotuberculosis is identified by cell and colony morphology, Gram stain, and reactions in biochemical and other tests. The bacterium is motile at 25 C. Serologic Detection in the Host. Serologic tests are not used to detect Y. pseudotuberculosis infections in poultry.

Pasteurella multocida INTRODUCTION

stage, particularly when the infection is caused by organisms of low virulence. Finding dead birds may be the first sign of fowl cholera. Other typical signs are depression, diarrhea, ruffled feathers, increased respiratory rate, and cyanosis. Commonly observed lesions in birds dying of acute fowl cholera include passive hyperemia, hemorrhages, swollen liver, focal necrotic areas in the liver and spleen, and increased pericardial and peritoneal fluids. In general, the signs of chronic fowl cholera include swelling of affected tissues, such as joints and sternal bursae, and exudate from conjunctivae and turbinates. The focal lesions are generally characterized by fibrinosuppurative exudate and various degrees of necrosis and fibroplasia.

The disease caused by infection with Pasteurella multocida is usually called fowl cholera; however, the term avian cholera is frequently used when the disease occurs in wild birds. The name Pasteurella septica infection was sometimes used in older European literature. Fowl cholera is a common, widely distributed disease of major economic importance in the United States. The disease affects all species of birds. Among commercially raised birds, turkeys and Japanese quail are particularly susceptible. Outbreaks in wild waterfowl are common and frequently cause high mortality. Fowl cholera occurs as a primary disease that does not require predisposing factors, although predisposing factors may increase severity of outbreaks. Subclinical infections apparently do not exist in normal flocks, but normal-appearing birds that have survived outbreaks of the disease frequently remain infected and may serve as carriers.

SAMPLE COLLECTION

Pasteurella multocida can be isolated readily from the liver, bone marrow, and heart blood of birds that die of acute fowl cholera and usually from localized lesions of chronic cholera (5). Bone marrow and brain are recommended when specimens are not fresh or when contamination of tissues seems likely (25). To obtain specimens for microbiologic examination, the surface of the tissue is seared with a heated spatula, and a sterile cotton swab or wire loop is inserted through the seared surface. The specimen is transferred to an agar

CLINICAL DISEASE Fowl cholera may affect birds of any age, but it rarely occurs in commercially raised poultry of less than 8 wk of age. The infection often occurs as an acute septicemic disease with high morbidity and mortality (5). Chronic fowl cholera may follow the septicemic

12

Chapter 4

Pasteurellosis, Avibacteriosis, Gallibacteriosis, Riemerelloisis, And Pseudotuberculosis

medium and incubated at 37 C. Pasteurella multocida grows aerobically and anaerobically. Cultures of P. multocida are moderately stable, generally surviving storage or transportation if maintained in a humid environment, such as on agar slants in screw-capped tubes. Stab cultures in agar medium in screw-capped tubes generally survive for weeks. Long­ term storage is best accomplished using lyophilization.

strains of P. multocida. A gel-diffusion precipitin test (GDPT) is used for serotyping based on differences in somatic antigens (somatic serotyping) (9). Sixteen serotypes (1-16) have been reported (2); strains representing each of these 16 serotypes have been isolated from avian hosts. Frequently, antigens from a single strain react with more than one type of serum, resulting in serotypes such as 3,4 and 3, 4, 12. An indirect (passive) hemagglutination test is used for serogrouping based on differences in capsular antigens (capsular serogrouping) (3). Five capsular serogroups (A, B, D, E, and F) have been reported (20). Serogroup A, D, and F strains produce capsules containing mucopolysaccharides, and presumptive identification of these serogroups can be made using specific mucopolysaccharidases in a disk-diffusion test (16). Except for serogroup E, strains representing all serogroups have been isolated from avian hosts. Antisera used in determining somatic serotypes are prepared in chickens (9, 17). Such typing antisera are available from the National Veterinary Services Laboratories, Animal and Plant Health Inspection Service (APHIS), United States Department of Agriculture (USDA), Ames, Iowa. Laboratories wishing to obtain such sera should contact the APHIS-USDA Veterinarian-in-Charge in their state. Antigens for the GDPT are prepared from 18-to-24-hr growth of heavily seeded DSA in petri dishes. The cells from one dish are suspended in 1.0 ml of 0.85% NaCl, 0.02M phosphate, and 0.3% formalin solution, pH 7.0. The suspension is heated in a water bath at 100 C for 1 hr and centrifuged at 4000 x g for 30 min. The supernatant fluid is used for antigen. The agar gel consists of 0.9% Noble hgar (Difco, Detroit, Mich.) in 8.5% NaCl solution. Five milliliters of warm (46 C) melted agar is flooded onto a 25 x 75-mm microscope slide; wells, 4 mm in diameter and 6 mm from center to center, are cut. Antigen is placed in a well and antisera are placed in opposing wells. The slide is placed in a petri dish to prevent drying, and the results are recorded after 24 hr at 37 C. Antisera used in the indirect hemagglutination test for capsule serogrouping are prepared in Pasteurellα-free rabbits (18, 20). Preparation of high-titered sera requires repeated intravenous inoculations with formalin-killed capsulated organisms. Currently, antisera for capsule serogrouping are not available from commercial or government laboratories.

PREFERRED CULTURE MEDIA AND SUBSTRATES Dextrose starch agar (DSA), blood agar, or trypticase soy agar (Becton Dickinson Microbiology Systems, Sparks, Md.) are recommended for primary isolation of P. multocida. The likelihood of isolation may be improved by supplementing these media with 5% heat-inactivated serum. The organisms grow readily in tryptose or trypticase soy broth.

AGENT IDENTIFICATION Colony Morphology On DSA, 24-hr colonies are circular, 1-3 mm in diameter, smooth, transparent, glistening, and butyrous. Colonies on blood agar are similar to those on DSA but are grayish and less translucent. Observation of 24-hr colonies on DSA or other translucent agar using a dissection microscope and obliquely transmitted lighting (5) provides information on whether or not cells are capsulated. Colonies that are iridescent contain capsulated cells, whereas colonies that are noniridescent (blue or blue-gray) contain uncapsulated cells. Pasteurella multocida produces a distinctive odor when grown on agar media.

Cell Morphology Pasteurella multocida cells are typically rods of 0.2-0.4 χ 0.6-2.5 pm occurring singly or occasionally in pairs or short chains. When grown under unfavorable conditions or after repeated subculture, cells tend to become pleomorphic. Cells in tissues or exudate usually show bipolar staining with Giemsa or Wright’s stain. Capsules can be demonstrated using an indirect india-ink method of staining. A loopful of dilute bacterial specimen is mixed with a loopful of india ink on a microscope slide, a coverslip is applied, and the preparation is examined microscopically at a high magnification. Capsules appear as clear halos around the bacterial cells when examined in this manner.

SEROLOGIC DETECTION IN THE HOST

Commercially available enzyme-linked immunosorbent assay (ELISA) kits may be used to detect a serological response to P. multocida infection in poultry (IDEXX, Westbrook, Maine and Synbiotics, Gaithersbury, MD). ELISAs are used primarily to measure the serologic response following the use of inactivated P. multocida vaccines in poultry. Serologic tests are rarely used for diagnosis of fowl cholera.

Biochemical and Other Tests Evaluation of reactions in differential media is generally made after 2 days of incubation at 37 C and again after 5 days of incubation at room temperature (21 C), in case of delayed reactions. The carbohydrate broth media used for identification (Table 4.1) is phenol red broth base containing 1% of the carbohydrate substrate. For detection of hydrogen sulfide (H2S), a filter paper strip impregnated with lead acetate is suspended above modified H2S broth (10) during incubation. The presence of indole is indicated by development of a dark red color when a small amount of modified Kovac’s indole test reagent is added to a 24-hr culture consisting of 2% tryptose in 0.85% NaCl solution (7). Oxidase production is determined using Kovac’s oxidase-test reagent and indirect filter paper procedure (12). Fructose, galactose, glucose, and sucrose are fermented without gas production. Inositol, inulin, maltose, salicin, and rhamnose are not fermented. Indole and oxidase are almost always produced. Neither hemolysis of blood nor growth on MacConkey’s agar occur. Differential characteristics are listed in Table 4.1.

DIFFERENTIATION FROM CLOSELY RELATED AGENTS

Differentiation of fowl cholera from other diseases caused by Pasteurella, Avibacterium, Gallibacterium, Riemerella anatipestifer, and Yersinia pseudotuberculosis depends primarily upon isolation and differentiation of causative organisms. Differential physiologic reactions of these organisms are indicated in Table 4.1. Several other diseases that are not caused by closely related agents, such as infectious coryza, fowl typhoid, fowl plague, duck plague, and infectious synovitis may have histories, signs, or lesions similar to those of fowl cholera. Differentiation of fowl cholera from these diseases also depends upon isolation and identification of causative organisms.

Serologic Identification Serologic tests are rarely used for identification of P. multocida. However, these are commonly used for antigenic characterization of

13

John R. Glisson, Tirath S. Sandhu, and Charles L. Hofacre

Table 4.1. Differential characteristics of Pasteurella, Avibacterium, Gallibacterium, Riemerella, and Yersinia.

Test

P. multocida

A. gallinarum

G.anatis biovar haemolytica

R. anatipestifer

Arabinose

-U

-

-

-

Dextrin

Dulcitol

-u -u

Fructose

+

Y. pseudotuberculosis

+

+

-U

+

+

+

+

Galactose

+

+

+

+

Glucose

+

+

+

+

Glycerol

+u

+

+

+

Inositol

-

V

+u

-

Inulin

-

-

-

-

Lactose

u

+

+u -u

+

Mannitol

+

+

+

+

+

Mannose

+

+

Maltose

+

Melibiose Raffinose

Rhamnose

-u -u

+

V

-

-

+

+U

+

V . +

Salicin

-

-

Sorbitol

+u

Sucrose

+

V +

Trehalose

-u +u

+u

Xylose

Gelatin

-

-

-

+ -

+u +u

Hemolysis

-

-

+

H2S

+u

+u

+u

+ +U

+

-

V

Indole

+

-

-

-

Litmus milk

-

si. acid U

si. alk

si. Alk

MacConkey

-

V

-

+U

-

+(25 C) +

Motility

-

-

Methyl red

Nitrate reduced

+

+

+

Oxidase

+

+

Urease

-

-

-

+

+

+

-

V

+

+

A+ = reaction, - = no reaction, U = usual reaction, V = variable reaction, si. Acid = slightly increased acidity, si. alk = slightly increased alkalinity, (25 C) =- incubated at 25 C.

Avibacterium gallinarum INTRODUCTION Swollen and inflamed wattles resulting from infection with only A. gallinarum have been reported in chickens (13).

Infection with A. gallinarum often occurs in the respiratoiy tracts of chickens and turkeys (6, 13), but disease is generally manifested only when this infection occurs with other respiratoiy tract infections. Attempts to isolate A.gallinarum from normal flocks have not been successful. The infection is widely distributed. It has been reported in the United States, Australia, Japan, Nigeria, Iran, and Israel. Generally it is not considered to be of economic significance.

SAMPLE COLLECTION

Avibacterium gallinarum often can be isolated from the sinuses, nasal cleft, trachea, air sacs, and lungs of birds exhibiting respiratory disease. The organism is less frequently isolated from livers, heart blood, and joints. Avibacterium gallinarum grows aerobically and anaerobically. Specimens are collected using the same methods as described for P. multocida.

CLINICAL DISEASE Disease manifestations in which A. gallinarum infection occurs usually affect the respiratory tract and have a chronic course.

PREFERRED CULTURE MEDIA AND SUBSTRATES

The same media are used as described for P. multocida. 14

Chapter 4

Pasteurella, Avibacteriosis, Gallibacteriosis, Riemerellosis, and Pseudotuberculosis

AGENT IDENTIFICATION

gallinarum using serologic tests, but specific serotypes have not been defined. Serologic testing has indicated that minor antigens of some strains of A. gallinarum are shared with P. multocida.

Colony and Cell Morphology and Biochemistry On DSA or blood agar at 37 C, 24-hr colonies are 1-2 mm in diameter, smooth, entire, low convex, and transparent. Colonies on DSA are iridescent when observed with obliquely transmitted light, often with concentric rings. Avibacterium gallinarum cells are similar to P. multocida; that is, Gram-negative short rods that occur as single organisms and short chains, stain bipolarly, and are capsulated. They become pleomorphic with repeated subculturing and growth under less than optimum conditions. Glucose, sucrose, and maltose are fermented without gas production. Lactose is not fermented, and indole is not produced. Neither hemolysis of blood nor growth on MacConkey’s agar occur. Differential characteristics are listed in Table 4.1.

SEROLOGIC DETECTION IN THE HOST

Serologic tests are not used to detect A. gallinarum infections in poultry. DIFFERENTIATION FROM CLOSELY RELATED AGENTS

Differentiation of A. gallinarum infections from those caused by Pasteurella and Y. pseudotuberculosis is accomplished by determination and comparison of selected physiologic characteristics (Table 4.1). Other infections that may cause similar signs or lesions include infectious coryza, mycoplasmosis, and bordetellosis. Differentiation depends upon isolation and identification or differentiation of the agents.

Serologic Identification Serologic tests are rarely used for identification of A. gallinarum. Antigenic differences have been demonstrated among strains of A.

Gallibacterium anatis biovar haemolytica INTRODUCTION

AGENT IDENTIFICATION

Gallibacterium anatis biovar haemolytica is generally considered to be a secondary pathogen that requires some predisposing factor before producing disease. However, too little information is available to clearly establish the role of this organism in avian disease production. In normal-appearing flocks, the reported incidence of infection is quite high in chickens but low in turkeys and geese. Infections in avian hosts are widely distributed, as indicated by isolations in the United States, England, Germany, France, Israel, Syria, Nigeria, Taiwan, and Australia. This disease is not considered economically important.

Colony Morphology On blood agar at 37 C, 24-hr colonies are generally 0.5 mm in diameter, smooth, entire, low convex, and translucent. A wide zone of β-hemolysis is produced on blood agar. On DSA, the colonies are iridescent when observed with obliquely transmitted light and often exhibit concentric rings.

Cell Morphology G. anatis biovar haemolytica cells are Gram-negative, nonmotile, non-spore-forming, pleomorphic rods (0.5 x 1.6 pm) that often exhibit bipolar staining. They generally occur singly but occasionally form short chains.

CLINICAL DISEASE

Disease manifestations related to G. anatis biovar haemolytica infections most frequently involve the respiratory tract. Septicemic manifestations with petechial hemorrhages in the viscera and areas of liver necrosis have been reported. Salpingitis and peritonitis may occur in laying hens.

Biochemical and Other Tests Glucose, sucrose, mannose, glycerol, and fructose are fermented without gas production within 24 hr of incubation at 37 C. Lactose is usually fermented after 3 days. Arabinose, dulcitol, rhamnose, inulin, and salicin are not fermented. Indole is not produced. Differential characteristics are indicated in Table 4.1.

SAMPLE COLLECTION Serologic Identification Serologic tests are not used for identification of G. anatis biovar haemolytica. Little effort has been made to serotype strains isolated from avian hosts, and no avian serotypes have been established.

G. anatis biovar haemolytica usually can be isolated from the trachea, lungs, liver, or oviduct of an infected bird.

SEROLOGIC DETECTION IN THE HOST Serologic tests are not used to detect G. anatis biovar haemolytica infection in poultry.

Riemerella anatipestifer the disease can cause mortality as high as 75% in ducks, especially at farms where infection persists because hatches are frequently moved from one pen to another to create space for the next hatch. Adverse environmental conditions and concomitant disease often predispose flocks to epomitics of R. anatipestifer infection. The disease is not of public health importance. In the United States, federal or state notification is not required.

INTRODUCTION

Riemerella anatipestifer infection, also known as infectious serositis, duck septicemia, new duck disease, or anatipestifer syndrome is a septicemic disease of ducks, geese, turkeys, and various other birds caused by R. anatipestifer. The disease is prevalent worldwide and causes significant economic loss due to high mortality, weight loss, and condemnations. The acute form of 15

John R. Glisson, Tirath S. Sandhu, and Charles L. Hofacre

long and 0.1-0.4 pm wide and may show bipolar staining. A capsule has been shown with the india ink method of staining. Riemerella anatipestifer does not ferment sugars in routine media (Table 4.1), but has been reported to produce acid in glucose, mannose, maltose and dextrin when grown in buffered single substrate medium (11). The organism liquefies gelatin and produces a slight alkaline reaction in litmus milk. Some strains produce urease and arginine dihydrolase. No growth occurs on MacConkey’s agar and no hemolysis takes place on blood agar. The organism produces oxidase, catalase, and phosphatase, but is indole negative. Riemerella anatipestifer is identified by enzymatic activity (Apizyme - API laboratory Products Ltd., St-Laurent, Quebec, Canada). It is positive for leucine-, valine-, and cystine-arylmidases, phosphoamidase, α-glucosidase, and negative for a- and βgalactosidases, β-glucuronidase, β-glucosidase, α-mannosidase, βglucosaminidase, ornithine and lysine decorboxylases (22).

CLINICAL DISEASE

The acute form of the disease usually occurs in ducklings 1-8 wk of age. Chronic infections may occur in older birds. High mortality has been reported in turkeys 6-15 wk of age (23). Riemerella anatipestifer infections have also been reported in swans, pheasants, guinea fowl, partridges, quail, and chickens. Clinical signs of the disease include ocular and nasal discharge, sneezing, greenish diarrhea, tremors of the head, neck and legs, ataxia, and coma. The common gross lesions are fibrinous pericarditis, perihepatitis, airsacculitis, and meningitis. In females, the oviduct is filled with caseous yellowish white exudate. Chronic and localized infections result in synovitis/arthritis and dermatitis. Infections originate from exposure via the respiratory tract or through abrasions or cuts in the skin. SAMPLE COLLECTION Riemerella anatipestifer can readily be isolated from heart blood, brain, pericardial exudate, air sacs, lungs, oviduct, and liver. Isolations should be attempted from infraorbital sinuses and trachea to detect carriers or inapparent infections. Specimens should be obtained by a wire loop or sterile swab through a seared surface. In chronic or localized infections, R. anatipestifer can be isolated from the exudate. The specimen is streaked directly onto the surface of agar media, which are then incubated at 37 C in a candle jar for 2472 hr. Incubation under increased CO2 and humidity in a candle jar or CO2 incubator enhances the growth for primary isolation. If the cultures cannot be made within a reasonable time, the specimens or tissues should be kept at 4 C or frozen for shipping or later processing.

Serologic Identification Using agglutination tests, 21 serotypes have been reported (15, 21). Antisera are made in rabbits. Antigen for rabbit inoculation is made by harvesting 24 to 48-hr growth from an agar plate in 0.85% NaCl solution containing 0.3% formalin. The inactivated cells are washed twice by centrifugation in the above solution and adjusted to an optical density of 0.2 at 525 nm in a spectrophotometer. Young rabbits are immunized by inoculating through the marginal ear vein successive doses of 0.05, 0.1, 0.2, 0.5, 1.0, 1.0, 1.5, and 2.0 ml of standardized cell suspension, at 3- to 4-day intervals. Rabbits are bled for serum collection when maximum titers are obtained. Agglutination titers are low, usually on the order of 1:50 to 1:400. The plate agglutination test is done by mixing a drop of undiluted antiserum with one or two colonies from a 24-hr growth. Agglutination within a few seconds indicates a homologous serotype reaction. The tube agglutination test is performed by mixing an equal amount of formalinized cell suspension (adjusted to 0.2 optical density at 525 nm) with each dilution of serum and incubating at 37 C. Agglutination is recorded after 24 and 48 hr.The fluorescent antibody technique may be used to detect and identify R. anatipestifer cells directly in tissues or exudate from infected birds.

PREFERRED CULTURE MEDIA AND SUBSTRATES

Although R. anatipestifer grows readily on routine media, growth is enhanced by using enriched media such as blood agar or trypticase soy agar containing 0.05% yeast extract. Tryptose broth or trypticase soy broth is used for broth cultures. Addition of gentamicin (5 mg/liter) to enriched solid medium has proven helpful for isolation of R. anatipestifer from sinuses, skin, and contaminated specimens from the processing plant, even though growth of other organisms is not completely inhibited.

SEROLOGIC DETECTION IN THE HOST

Enzyme-linked immunosorbent assay (ELISA) is commonly used for early detection of R. anatipestifer infections (8). Cell lysate is used as an antigen in ELISA. It is not serotype specific and will show positive reactions with antisera against heterologous serotypes, but ELISA is more sensitive than agglutination test. The GDPT is unreliable because of an apparent discrepancy in duck immunoglobulins, which are almost nonfunctional in precipitin reactions (24).

AGENT IDENTIFICATION Colony and Cell Morphology and Biochemistry Riemerella anatipestifer is identified on the basis of growth, morphologic and biochemical characteristics. After 24 hr of growth on blood agar, colonies are 1-2 mm in diameter, convex, entire, transparent, glistening, and butyrous. Some strains may produce mucoid growth on solid media. On trypticase soy agar, colonies appear bluish and are iridescent when observed with obliquely transmitted light. Some strains have been observed to produce offwhite growth, which may change to grayish brown after 3-5 days. Growth in broth produces slight turbidity. Most R. anatipestifer strains become nonviable after 3^4 days on a solid medium at room temperature or 37 C. In broth the organisms may survive for 2-3 wk under refrigeration. Cultures can be stored as freeze-dried for longer periods. Riemerella anatipestifer cells are Gram-negative, nonmotile, non-spore-forming rods that occur singly, in pairs, or occasionally in chains. The cells are 1-5 pm

DIFFERENTIATION FROM CLOSELY RELATED AGENTS

Diagnosis should be based on isolation and identification of the causative organism, because similar gross lesions are produced by P. multocida, Escherichia coli, and fecal streptococcal infections. Ducklings infected with Salmonella may sometime show nervous signs similar to those of R. anatipestifer infection. Chlamydiosis should also be considered in differential diagnosis.

16

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Pasteurella, Avibacteriosis, Gallibacteriosis, Riemerellosis, and Pseudotuberculosis

Yersinia pseudotuberculosis INTRODUCTION

Cell Morphology Yersinia pseudotuberculosis cells are Gram-negative rods of 0.5 x 0.8-5.0 pm. Coccoid forms usually stain bipolar. Neither spores nor capsules are formed. Peritrichous flagella usually develop between 20 C and 30 C. Motility is best shown in semisolid media at 25 C.

The disease caused by infection with Y.pseudotuberculosis is pseudotuberculosis (19). It can affect domesticated, feral, or caged birds and a variety of mammals, including humans. Although this disease is infrequently reported in the United States and is not considered economically important, it has caused death losses as high as 80% in turkey flocks.

Biochemical and Other Tests Maltose, trehalose, and usually sucrose are fermented, and acid but not gas is produced. Production of H2S is variable. Urease and catalase are produced, but not indole and oxidase. No hemolysis occurs on blood agar. Differential characteristics are listed in Table 4.1. Serologic Identification Serologic testing is not used for identifying strains of Y. pseudotuberculosis but is used to characterize strains (19). There are six serotypes (I-VI), four of which have two subtypes each (A and B). These serotypes and subtypes are based on 15 heat-stable somatic antigens (1-15), which are demonstrated by agglutination and agglutination-adsorption tests. Additional serotypes VH and Vin and a subtype C of serotype Π have been proposed. Among strains from birds, serotype I is most common, followed by serotypes Π and IV. Serotypes V and VI have not been reported in birds, and type ΠΙ is rare. Five heat-labile flagellar antigens (a, b, c, d, e) are recognized but are rarely used for serologic characterization.

CLINICAL DISEASE

Pseudotuberculosis usually occurs as a septicemia or bacteremia of short duration, followed by chronic focal infections. In very acute cases, birds may die before other signs of disease are observed, although signs usually are present for 2 wk or more. Affected birds exhibit weakness, ruffled feathers, diarrhea, and breathing difficulties. Emaciation and paralysis occur occasionally. Swollen livers and spleens and enteritis are common lesions in acute cases. Necrotic foci in visceral organs and muscles and enteritis are common lesions in chronic cases. Osteomyelitis occurs in some affected turkey flocks. SAMPLE COLLECTION Yersinia pseudotuberculosis can usually be isolated from blood, liver, spleen, or lung using blood or trypticase soy agar and incubation at 37 C (19). The method of Paterson and Cook (14) is recommended for isolation of Y pseudotuberculosis from feces. To use this method, inoculate the surface of Paterson and Cook agar medium with a 10% suspension of feces in phosphate buffer (pH 7.6). Recovery of Y. pseudotuberculosis is enhanced by storing the fecal suspension at 3-4 C for 7 days or longer before inoculating agar. The organism grows both aerobically and anaerobically.

SEROLOGIC DETECTION IN THE HOST Serologic techniques are not used to detect Y. pseudotuberculosis infection in poultry. DIFFERENTIATION FROM CLOSELY RELATED AGENTS

Trypticase soy agar and blood agar are effective for primary isolation. Differential media are as described for P. multocida. For isolation from feces, Paterson and Cook medium is recommended (14) (Table 4.2).

Differentiation of Y. pseudotuberculosis from avian Pasteurella is based on differences in biochemical characteristics, as indicated in Table 4.1. Diseases with similar lesions or signs, such as tuberculosis, E. coli infections, and salmonellosis, are differentiated from pseudotuberculosis through isolation and identification or biochemical differentiation of causative agents.

AGENT IDENTIFICATION

REFERENCES

Colony Morphology After incubation at 37 C for 24 hr on trypticase soy agar, colonies are smooth, round, entire, grayish yellow, butyrous, and 0.5-1 mm in diameter. Older colonies are 2-3 mm in diameter, raised, flat, dry, and irregular with rough edges.

1. Blackall, P.J. H. Christensen, T. Beckenham, L.L. Blackall, and M Bisgaard Reclassification of Pasteurella gallinarum, [Haemophilus] paragallinarum, Pasteurella avium and Pasteurella volantium as Avibacterium gallinarum gen. nov., comb, nov., Avibacterium paragallinarum comb, nov., Avibacterium avium comb, nov. and Avibacterium volantium comb. nov. Int. J. Evol. Microbiol. 55: 353-362, 2005. 2. Brogden, K. A., K. R. Rhoades, and K. L. Heddleston. A new serotype of Pasteurella multocida associated with fowl cholera. Avian Dis. 22:185190. 1978. 3. Carter, G. R. Studies on Pasteurella multocida. I. A hemagglutination test for the identification of serological types. Am. J. Vet. Res. 16:481^484. 1955. 4. Christensen, Η., M. Bisgaard, A.M Bojesen, R. Mutters, and J.E. Olsen. Genetic relationships among avian isolates classified as Pasteurella haemolytica, Actinobacillus salpingitidis or Pasteurella anatis with proposal of Gallibacterium anatis gen. nov., comb. nov. and description of additional genomospecies with Gallibacterium gen. nov. Int. J. Syst. Evol. Microbiol. 53: 275-287,2003. 5. Glisson, J. R., C. L. Hofacre and J. P. Christensen. Fowl cholera. In: Diseases of poultry, 11th ed Y. M. Saif, H. J. Barnes, J. R. Glisson, A M Fadly, L. R. McDougald, and D. E. Swayne, eds. Iowa State University Press, A Blackwell Publishing Company, Ames, Iowa. pp. 658-676. 2003.

PREFERRED CULTURE MEDIA AND SUBSTRATES

Table 4.2. Medium of Paterson and Cook. Constituents Trypsinized meat agar

Peptic digest of sheep blood Novobiocin (0.2%)

Erythromycin (0.2%) Mycostatin (50,000 units/ml) Crystal violet (0.1%)

Amount (ml) 200.0

10.0 0.5 0.5

0.8 0.5

17

John R Glisson, Tirath S. Sandhu, and Charles L. Hofacre

6. Hall, W. J., K. L. Heddleston, D. H. Legenhausen, and R. W. Hughes. Studies on pasteurellosis. I. A new species of Pasteurella encountered in chronic fowl cholera. Am. J. Vet. Res. 16:598-604. 1955. 7. Hans, G. H., and S. Gabriel. Modified stable Kovac’s reagent for detection of indole. Am. J. Clin. Pathol. 26:1373-1375. 1956. 8. Hatfield, R. Μ., B. A Morris, and R. R. Henry. Development of an enzyme-linked immunosorbent assay for the detection of humoral antibody to Pasteurella anatipestifer. Avian Pathol. 16:123-140. 1987. 9. Heddleston, K. L., J. E. Gallagher, and P. A. Rebers. Fowl cholera: gel difiusion precipitin test for serotyping Pasteurella multocida from avian species. Avian Dis. 16:925-936. 1972. 10. Heddleston, K. L., T. Goodson, L. Liebovitz, and C. I. Angstrom. Serological and biochemical characteristics of Pasteurella multocida from free-flying birds and poultry. Avian Dis. 16:925-936. 1972. 11. Hinz, K. Η., M Ryll, and B. Kohler. Detection of acid production from carbohydrates by Rimerella anatipestifer and related organisms using the buffer^ single substrate test. Vet. Microbiol. 60:277-284. 1998. 12. MacFaddin, J. F. Oxidase test. In: Biochemical tests for identification of medical bacteria, 2nd ed. Williams and Wilkins, Baltimore, Md. pp. 249-253. 1980. 13. Mushin, R., R. Bock, and M Abrams. Studies on Pasteurella gallinarum. Avian Pathol. 6:415-423. 1977. 14. Paterson, J. S. and R Cook. A method for the recovery of Pasteurella pseudotuberculosis from feces. J. Pathol. Bacteriol. 85:241-242. 1963. 15. Pathanasophon, P., P. Phuektes, T. Tanticharoenyos, W. Narongsak, and T. Sawada. A potential new serotype of Riemerella anatipestifer isolated from ducks in Thailand. Avian Pathol. 31:267-270. 2002. 16. Rimler, R. B. Presumptive identification of Pasteurella multocida serogroups A, D, and F by capsule depolymerisation with mucopolysaccharidases. Vet. Rec. 134:191-192. 1994. 17. Rimler, R B., R. D. Angus, and M Phillips. Evaluation of the specificity of Pasteurella multocida somatic antigen-typing antisera prepared in chickens using ribosome-lipopolysaccharide complexes as inocula. Am. J. Vet. Res. 50:29-31. 1989. 18. Rimler, R. B., and K. A. Brogden. Pasteurella multocida isolated from rabbits and swine: serologic types and toxin production. Am J. Vet. Res. 47:730-737. 1986.

19. Rimler, R B., and J. R. Glisson. Pseudotuberculosis. In: Diseases of poultry, 10th ed. B. W. Calnek, H. J. Bames, C. W. Beard, L. R. McDougald, and Y. M Saif, eds. Iowa State University Press, Ames, Iowa, pp. 314-317. 1997. 20. Rimler, R. B., and K. R. Rhoades. Serogroup F, a new capsule serogroup of Pasteurella multocida. J. Clin. Microbiol. 25:615-618. 1987. 21. Sandhu, T. S. Riemerella anatipestifer infection. In: Diseases of poultry, 11th ed. Y. M Saif, H. J. Bames, J. R. Glisson, A. M Fadly, L. R. McDougald, and D. E. Swayne, eds. Iowa State University Press, A Blackwell Publishing Company Ames, Iowa. pp. 676-682. 2003. 22. Segers, P., W. Mannheim, M Vancanneyt, K. DeBrandt, K. H. Hinz, K. Kersters, and P. Vandamme. Riemerella anatipestifer gen. nov., comb, nov., the causative agent of septicemia anserum exudativa, and its phylogenetic affiliation within the Flavobacterium-Cytophaga rRNA homology group. Int. J. Syst. Bacteriol. 43:768-776. 1993. 23. Smith, J. M, D. D. Frame, G. Cooper, A. A. Bickford, G. Y. Ghazikhanian, and B. J. Kelly. Pasteurella anatipestifer infection in commercial meat-type turkeys in California. Avian Dis. 31:913-917. 1987. 24. Toth, T. E., and N. L. Norcross. Precipitating and agglutinating activity in duck anti-soluble protein immune sera. Avian Dis. 25:338-352. 1981. 25. Waltman, W.D. and A.M Home. Characteristics of fowl cholera diagnosed in Georgia 1989-1991. Avian Dis. 37:616-621, 1993.

18

5 BORDETELLOSIS Mark W. Jackwood SUMMARY. Bordetellosis is an acute highly contagious disease of the upper respiratory tract of young turkeys (4-8 wk of age). The disease is caused by a gram-negative nonfermentative bacteria, Bordetella avium. Members of the genus Bordetella are well known for their ability to colonize and damage ciliated epithelial surfaces in the respiratory tract and B. avium is no exception. Bordetellosis is most severe when it occurs in conjunction with other respiratory infections, especially Newcastle disease, or when turkeys are immunosuppressed. Mortality can be high when poor management (inadequate ventilation, dust, chilling, or filthy conditions) is a complicating factor and there is an associated colibacillosis. Bordetella avium is an opportunistic pathogen in chickens. Agent Identification. Bordetellosis can be diagnosed using a combination of clinical signs (snick or cough with a catarrhal nasal discharge) and bacterial culture. Serologic Detection in the Host. Enzyme-linked immunosorbent assays and microagglutination tests are accurate and sensitive for detecting antibodies in turkeys. Test results can be an aid in diagnosis as well as monitoring vaccination.

sp. Cultures should be taken early during the infection while cilia are heavily colonized with B. avium. If cultures are taken late in the infection the most predominant bacteria isolated will likely be Escherichia coli.

INTRODUCTION In the 1970s an acute upper respiratory disease emerged as a major disease problem in young turkeys. The disease was more severe the earlier posthatch turkeys become infected. Initially some confusion occurred as to the identification of the causative agent but a definitive study established the agent as a new species in the genus Bordetella and it was given the name Bordetella avium (7). In the early years following its recognition, the disease and associated colibacillosis produced high mortality but in recent years the disease or the susceptibility of turkeys to the disease has changed and the number and severity of reported outbreaks has lessened. Some of the possible explanations for the change in the character of the disease include the presence of maternal antibody, as it is well established that passive immunity is highly protective (9), or that strains of B. avium have emerged that are not as virulent. The significance of bordetellosis is not as great now as when it was first recognized but it still is considered to be a major cause of respiratory disease in young turkeys. Bordetella avium has also been isolated from chickens. Bordetella avium is an opportunistic pathogen in chickens because clinical disease can only be reproduced following initial infection with an upper respiratory disease virus vaccine such as infectious bronchitis virus or Newcastle disease virus (4). Recent studies have reported B. avium in mallards, wild turkeys, and a Canada goose, (10) and antibodies against B. avium have been detected in peafowl (2).

PREFERRED CULTURE MEDIA AND SUBSTRATES Bordetella avium grows on a variety of enriched agar-based media but the bacteria is usually isolated from clinical material using MacConkey agar because this agar quickly demonstrates the bacteria’s nonfermenting characteristic and differentiates it from E. coli, another common bacteria found in the trachea of turkeys with respiratory disease. Initial isolation of B. avium is best done on MacConkey agar increased to 2.5% agar content to slow the spread of faster growing organisms (6). Growth of B. avium in liquid media requires that the cultures be aerated by agitation or other means because it is a strict aerobe. AGENT IDENTIFICATION

After 24 hr of incubation on MacConkey agar, colonies of B. avium are clear and pinpoint in size. If cultures have been secured early they are often pure but when cultures are taken late there is often contamination with other bacteria especially E. coli. When such contamination occurs it is important to look in the more diluted part of the streak for typical colonies. Three colony types of B. avium have been identified. The most typical type is that described above with colonies being small, compact, translucent, and pearl-like with entire edges and glistening surfaces. These colonies will be 0.2-1 mm in diameter after 24 hr and 1-2 mm after 48 hr. Many isolates develop a raised browntinged center when grown to >48 hr on MacConkey’s agar. A smaller percentage of isolates dissociate into a rough colony type having a dry appearance and a serrated irregular edge. Rough colonies represent a phase-shift to a nonpathogenic form of the bacterium (3). Biochemical tests that are used in distinguishing B. avium from other nonfermenting bacteria include the oxidase test (+), catalase test (+), urease test (-), nitrate reduction test (-), and the ability to alkalinize certain amides and organic salts. A microcupule test kit called the Rapid API 20 NE (Non-Fermenter Test, bioMerieuxVitek, Hazelwood, Mo.) can be used to identify B. avium. A very useful correlation has been made between the ability of strains of B. avium to hemagglutinate guinea pig red blood cells and pathogenicity (6). This association with pathogenicity is thought to be related to the ability of pathogenic strains to adhere to cilia. The hemagglutination assay is performed by heavily inoculating solid media (blood agar, brain-heart infusion agar, or MacConkey’s agar) with suspected B. avium isolates and incubating

CLINICAL DISEASE

Bordetellosis is characterized by an abrupt onset of a snick (sneezing) in young turkeys. Other signs include watery eyes, submaxillary edema, and a clear nasal discharge that can usually be expressed by placing gentle pressure on the nares. Mouth breathing, altered vocalization, dyspnea, huddling, and decreased consumption of feed and water are also common. Morbidity is high 80-100% in young turkeys, and mortality is low (1% to 10%) except when other infections occur simultaneously (colibacillosis) or when management is poor, then mortality can be high (>50%). The infection also adversely affects growth rate and toxins associated with the bacteria damage tracheal cartilage beneath colonized ciliated respiratory epithelium. Mortality has been reported from suffocation due to excessive mucus production in the trachea and when damaged tracheas collapse (12).

SAMPLE COLLECTION The bacterium is best cultured by swabbing the anterior trachea through a midcervical aseptic opening. Bacterial culture of sinus material is usually not productive because of overgrowth by Proteus

19

Mark W. Jackwood

36-48 hr at 37 C. Bacterial growths are washed from the plate with phosphate-buffered saline (PBS) and diluted to a concentration of approximately 5 χ 109 cells/ml (0.5 optical density at 600 nm). Equal volumes of bacterial suspension and erythrocyte suspension (2% packed-cell volume in PBS) are mixed on a glass slide or plate and observed for agglutination after gentle rocking. Alternatively, 50:1 mixture of bacterial suspension and erythrocyte suspension (0.5% packed-cell volume in PBS) are mixed in U-bottom shaped wells of microtitration plates and allow to stand 1 hr at 25 C. A reference strain of B. avium and a negative control should be included with each test run. Comparable results can be obtained using erythrocytes fixed in formaldehyde (F. W. Pierson, Virginia Polytechnic Institute and State University, pers. comm.). Fixed guinea pig erythrocytes are prepared as follows: 1. Collect 5 ml of guinea pig blood with 1 ml of 3% sodium citrate and mix well. 2. Centrifuge at 500 x g (2000 rpm) for 10 min. Resuspend erythrocytes in fixative containing 40 ml of PBS and 10 ml of 40% formaldehyde. The final pH of the solution should be approximately 7.4. 3. Mix gently on a rotary shaker at room temperature overnight. 4. Centrifuge at 500 x g (2000 rpm) for 10 min and wash pellet 10 times with PBS. 5. Resuspend fixed erythrocytes in PBS at a final concentration of 2% and store at 4 C. lite fixed cells are stable for at least 6 mos.

2. Drop 0.1 ml of stock antigen into row A, columns 1-8 of each plate. 3 Drop 0.05 ml of PBS with 0.01% merthiolate into rows B through H, columns 1-8. 4 Make twofold dilutions of B. avium antigen down from row A through row H. 5. Add 0.05 ml of the appropriate serum dilution (1:10 through 1:640, in columns 1 through 7, respectively) to rows A through H. 6. Add 0.05 ml PBS with 0.01% merthiolate to column 8, rows A through H, for the antigen control. 7. Incubate plates at room temperature for 24 hr and read. A positive test has complete absence of a button, whereas a negative test has a distinct button in the bottom of the well. For the optimum B. avium antigen dilution, pick the antigen dilution well that has a well-defined, visible button in the negative control serum and detects antibody (no button) in the positive control serum. It is necessary to do a block titration only once for each lot of B. avium antigen produced. Mark the appropriate dilution on the stock antigen for later reference. The actual microagglutination test is run as follows: 1. Make a 1:5 dilution of each serum sample, including both positive and negative controls. 2 Drop 0.05 ml PBS with 0.01% merthiolate into each well of a microtitration plate. 3. Drop 0.05 ml of diluted serum to the first row of wells. Be sure to include positive and negative control sera with each plate. 4. Make twofold dilutions of the sera from 1:10 to 1:1280 down the plate. 5. Dilute B. avium stock antigen in PBS with 0.01% merthiolate using the dilution factor from the block titration. The antigen should be passed through a syringe with a 22 gauge needle to break up all antigen clumps. 6. Drop 0.05 ml of diluted B. avium antigen into all wells. 7. Incubate the test plate at room temperature for 24 hr and read. Any titer 1:20 or greater is considered to be positive. The microagglutination test is not as user friendly as the ELISA but the test is sensitive and accurate. It detects immunoglobulin M (IgM), which is the first antibody produced following infection. The microagglutination test will not detect maternal antibody, which is immunoglobulin G (IgG). High levels of maternal antibody protect poults during the first 2 wk of life when the birds are most susceptible to the disease (12). Antibody titers (IgG) to B. avium can be detected by ELISA 2 wk postinfection, with peak titers occurring between 3 and 4 wk postinfection (5,8,9).

SEROLOGIC DETECTION IN THE HOST An in-house enzyme-linked immunosorbent assay (ELISA) can be produced to detect B. avium antibodies in turkeys (8,9). A commercially available ELISA has been developed and marketed for B. avium in turkeys (Synbiotics Corporation, San Diego, CA.). The commercially available ELISA kit is excellent and the results are very compatible with the in-house ELISA test (9). Prior to the development of the ELISA a microagglutination procedure was performed (5) as outlined below. Microagglutination antigen is produced as follows: 1. Inoculate 10 ml of veal infusion broth or brain-heart infusion broth with an isolated colony of B. avium and incubate aerobically at 37 C for 48 hr in a shaking-water bath or shaking incubator. 2. Inoculate 500 ml of sterile broth in a 1000-ml flask with 10 ml of broth culture from step 1. Incubate in a shaking-water bath or incubator at 37 C for 48 hr. When given ideal growth conditions, B. avium will sometimes produce a filamentous form in broth culture, which interferes with the microagglutination test (1). This problem can be circumvented by growing the bacterium on agar and harvesting the cells in PBS. Then, continue with step 3. 3. At 48, 49, and 50 hr, add 2.5 ml of a 1% solution of neotetrazolium chloride stain. Incubate 4 hr at 37 C following the last addition of neotetrazolium chloride. 4. Add 0.2 ml of a 0.1% solution of merthiolate and incubate overnight at 37 C. 5. Centrifuge for 30 min at 10,960 x g. Wash and centrifuge antigen three times in PBS with 0.01% merthiolate. 6. Centrifuge antigen in a graduated tube at 10,960 x g for 20 min, discard the supernatant, and resuspend to make a 1:20 dilution of packed cells in PBS with 0.01% merthiolate. This is the stock antigen. Pass the stock antigen through a 22-gauge needle with a syringe, run a block titration, aliquot the remaining antigen, and freeze the aliquots at -30 C or below. Microagglutination antigen standardization (block titration) is as follows: 1. Use separate microtitration plates for both B. avzwm-positive and negative sera. First make serial twofold dilutions of sera in test tubes beginning with 1:10 and ending with 1:640. About 0.5 ml of each dilution is needed for one microtitration plate.

DIFFERENTIATION FROM CLOSELY RELATED AGENTS

Bordetellosis in poultry should be differentiated from mycoplasmosis, chlamydiosis, cryptosporidiosis, Omithobacterium rhinotracheale, and viral respiratory diseases, particularly Newcastle disease, influenza, and turkey rhinotracheitis (pneumovirus infection). The most closely related agents to be differentiated are Bordetella bronchiseptica and Bordetella hinzii (also referred to as B. avium-tike bacteria in the literature). Both can be isolated from the upper respiratory tract, but neither have shown to cause disease in turkeys. B. avium causes agglutination of guinea pig erythrocytes, is urease-negative, and does not grow on minimal essential medium agar or in 6.5% NaCl broth. B. bronchiseptica may cause hemagglutination, but it produces a positive reaction for urease. B. hinzii does not hemagglutinate but will grow on minimal essential medium agar and in 6.5% NaCl broth (6). Molecular tests have been used to distinguish between B. avium and B. hinzii. Ribotyping using PvuH or restriction endonuclease analysis using Hinfl and DdeV enzymes clearly distinguish between the two organisms but the REA test was found to be best as a routine epidemiologic tool (11). 20

Chapter 5

Bordetellosis

REFERENCES

8. Lindsey, D. G., P. D. Andrews, G. S. Yarborough, J. K. Skeeles, B. Glidewell-Erickson, G. Campbell, and Μ B. Blankford. Evaluation of a commercial ELISA kit for detection and quantification of antibody against Bordetella avium. In: Proc. 75th Annual Meeting of the Conference of Research Workers in Animal Diseases, Chicago, Ill. Abstract #31. Nov. 14—15,1994. 9. Neighbor, N. K., J. K. Skeeles, J. N. Beasley, and D. L. Kreider. Use of an enzyme-linked immunosorbent assay to measure antibody levels in turkey breeder hens, eggs, and progeny following natural infection or immunization with a commercial Bordetella avium bacterin. Avian Dis. 35:315-320. 1991. 10. Raffel, T. R., K. B. Register, S. A. Marks, and L. Tempel. Prevalence of Bordetella avium infection in selected wild and domesticated birds in the eastern USA. J. Wildl. Dis. 38:40-46. 2002. 11. Register, K. B., R. E. Sacco, and G. E. Nordholm. Comparison of ribotyping and restriction enzyme analysis for inter- and intraspecies discrimination of Bordetella avium and Bordetella hinzii. J. Clin. Microbiol. 41:1512-1519. 2003. 12. Skeeles, J. K., and L. H. Arp. Bordetellosis (turkey coryza). In: Diseases of Poultry, 10th ed. B. W. Calnek, H. J. Barnes, C. W. Beard, L. R. McDougald, and Y. M Saif. Iowa State University Press, Ames, Iowa. pp. 275-287. 1997.

1. Domingo, D. D., M. W. Jackwood, and T. P. Brown. Filamentous forms of Bordetella avium', culture conditions and pathogenicity. Avian Dis. 36:706-713. 1992. 2. Hollamby, S., J. G. Sikarskie, and J. Stuht. Survey of peafowl (Pavo cristatus) for potential pathogens at three Michigan zoos. J. Zoo Wild. Med. 34:375-379. 2003. 3. Jackwood, M W., D. A. Hilt, S. M Mendes, and P. D. Cox. Bordetella avium phase-shift markers: characterization of whole cell, cell envelope, and outer membrane proteins. J. Vet. Diag. Invest. 7:402-404. 1995. 4. Jackwood, M. W., S. M McCarter, and T. P. Brown. Bordetella avium'. an opportunistic pathogen in leghorn chickens. Avian Dis. 39:360-367. 1995. 5. Jackwood, D. J., and Y. M Saif. Development and use of a microagglutination test to detect antibodies to Alcaligenes faecalis in turkeys. Avian Dis. 24:685-701. 1980. 6. Jackwood, M W.,Y. M. Saif, P. D. Moorhead, and R. N. Dearth. Further characterization of the agent causing coryza in turkeys. Avian Dis. 29:690-705. 1985. 7. Kersters, K., K.-H. Hinz, A. Hertle, P. Segers, A. Lievens, O. Siegmann, and J. De Ley. Bordetella avium sp. nov. isolated from the respiratory tracts of turkeys and other birds. Int. J. Syst. Bacteriol. 34:5670. 1984.

21

6 INFECTIOUS CORYZA Pat J. Blackall

SUMMARY. Infectious coryza, an acute upper respiratory tract disease of chickens, is caused by the bacterium Avibacterium paragallinarum. The main impact of the disease is a drop in egg production. Other manifestations such as airsacculitis in broilers, swollen head-like syndrome, arthritis, and septicemia are either unusual or probably due to complications associated with other infectious agents. Agent Identification. Diagnosis of infectious coryza is preferably made by the isolation and identification, by biochemical properties, of the bacterium. A polymerase chain reaction (PCR) test, which can be applied either to suspect colonies or directly to samples from chickens, is now available. The PCR test is particularly suited for those laboratories that lack suitable expertise and experience in the growth and phenotypic identification of A. paragallinarum or where samples are transported for long periods to the laboratory. Although most isolates of A. paragallinarum are dependent upon V factor for growth in artificial media (meaning they show the traditional satellitic growth), some isolates are V factor-independent. This variation in growth factor requirements, along with the existence of nonpathogenic V factor­ dependent organisms, increases the need for biochemical identification or the use of the PCR test. Serotyping of isolates is important to guide the use of vaccines. Serologic Detection in the Host. A range of serologic tests to detect antibodies has been described with hemagglutination-inhibition tests now being the most widely used.

When the disease occurs in chicken flocks in developing countries, the added presence of other pathogens and stress factors can result in disease outbreaks that are associated with greater economic losses than those reported in high health flocks in developed countries. In China, outbreaks of infectious coryza have been associated with morbidities of 20 to 50% and mortalities of 5 to 20% (13). In India, outbreaks of infectious coryza are often complicated with fowl cholera and can result in high mortalities e.g. one farm experienced 50% mortality in a 14,000 bird layer flock (33). Arthritis and septicemia, possibly complicated by the presence of other pathogens, have been reported in broiler and layer flocks in South America (27). Infectious coryza can also be a problem in the village production system e.g. there are reports of outbreaks in such chickens in Thailand (32) and Indonesia (24). Overall, there is considerable evidence that infectious coryza outbreaks can have a much greater impact in developing countries than in developed countries.

INTRODUCTION

Infectious coryza is an acute respiratory disease of chickens. The clinical syndrome has been recognized since the 1930s and has been described in the early literature as roup, contagious or infectious catarrh, and uncomplicated coryza (6). Early workers identified the causative agent as Haemophilus gallinarum, an organism that required both X (hemin) and V (nicotinamide adenine dinucleotide; NAD) factors for growth in vitro. However, from the 1960s to the 1980s, all isolates of the disease-producing agent have been shown to require only V factor and have been termed Haemophilus paragallinarum (6). V factor-independent H. paragallinarum isolates have been encountered in the Republic of South Africa (21) and Mexico (15). Most recently, a polyphasic taxonomic study has shown that the Haemophilus paragallinarum is not a member of the genus Haemophilus (2). Thus, H. paragallinarum was allocated to a new genus - Avibacterium - along with several other chicken associated members of the family Pasteurellaceae (2). Hence, the causal agent of infectious coryza is now called Avibacterium paragallinarum, an organism that can be either V factor-dependent or -independent. This text will use the new terminology of A. paragallinarum - even if the primary publications being cited used the older terminologies. The disease occurs worldwide and causes economic losses due to an increased number of culls and a marked reduction from 10% to more than 40% in egg production, particularly on multiage farms. The disease is essentially limited to chickens and does not threaten public health.

SAMPLE COLLECTION Two to three acutely diseased chickens should be killed and the skin over their sinuses seared with a heated spatula. The skin is then incised with a sterile scalpel blade, and a sterile cotton swab is inserted into the sinus cavity. Typically, in the early phase of the disease, A. paragallinarum is found in pure culture in the sinus. Swabs of the trachea and air sacs may be taken, although the organism is less frequently isolated from these areas. Avibacterium paragallinarum is a fragile organism that does not survive for more than 5 hr outside of birds. Sinus swabs held in Ames Transport medium (without charcoal) can yield positive cultures for up to 8 days at transport temperatures of either 25 C or room temperature (9). Live bird sampling is also possible. In this technique, gentle milking pressure is exerted on the sinus area and mucus forced from the nostril. The expressed mucus should be sampled by a bacteriologic loop, with care being taken to avoid touching the surface of beak or nostril. A swab of the expressed mucus is also the optimal sample for the A. paragallinarum PCR. Swabs collected by this method will still yield a PCR positive reaction after storage in glycerol-enriched phosphate buffered saline for 180 days at either 4 C or -20 C (12).

CLINICAL DISEASE

Infectious coryza may occur in growing chickens and layers. The most common clinical signs are nasal discharge, facial swelling, lacrimation, anorexia, and diarrhea. Decreased feed and water consumption retards growth in young stock and reduces egg production in laying flocks (6). In layers, the disease can have a modi greater impact than the relatively simple scenario described above. As an example, an outbreak of the disease in older layer birds in California, which was not associated with any other pathogen, caused a total mortality of 48% and a drop in egg production from 75 to 15.7% over a 3 wk period (8). The disease can have significant impact in meat chickens. In California, two cases of infectious coryza, one complicated by the presence of Mycoplasma synoviae, caused a swollen head like syndrome and increased condemnations, mainly due to airsacculitis, that varied from 8.0 to 15% (14).

PREFERRED CULTURE MEDIA AND SUBSTRATES

Artificial Media Blood agar is commonly used for the isolation of A. 22

Chapter 6

paragallinarum. The medium is prepared from a dehydrated base such as Bacto-tryptose-blood-agar base (Difco, Detroit, Mich.) and is enriched with 5% erythrocytes, which may be from any animal. The inoculum is streaked onto the blood agar plate in the conventional maimer, after which the plate is cross-streaked with a nurse culture. Blood agar is deficient in V factor, and the role of the nurse culture is to excrete excess V factor to support the growth of A. paragallinarum . Although several bacterial species are possible nurse cultures, it is recommended that Staphylococcus hyicus, a normal inhabitant of the skin of chickens, be used. Avibacterium paragallinarum is typically grown in the presence of 5% CO2 at 37 C. A convenient procedure is to use candle jars if CO2 incubators are not available. An alternative isolation (and maintenance) medium that is particularly suited to laboratories in developing countries has been described by Terzolo et al. (31). This medium consists of Columbia blood agar base (Becton Dickinson Microbiology Systems, Sparks, Md.) with 7% lysed equine blood. The lysed equine blood is prepared by holding fresh equine blood at 56 C for 40 min with occasional stirring. The lysed blood can be stored at -20 C. A selective version of the medium, which should only be used in parallel with the nonselective medium, contains bacitracin (5 U/ml), cloxacillin (5 pg/ml), and vancomycin (25 pg/ml). The plates are incubated under a microaerophilic atmosphere. Several complex media have been described that support growth of avian hemophili (a term used generically to refer to bacteria within this group). Such media, although not suitable for isolation due to problems with overgrowth by contaminants, are particularly useful for characterization tests following initial isolation. Two media that have proven very useful are Haemophilus maintenance medium (HMM) and supplemented test medium agar (TM/SN), originally described by Rimler et al. (25, 26). HMM base consists of 1% polypeptone (BBL), 1% biosate peptone (BBL), 0.24% beef extract, 0.005% para-aminobenzoic acid, 0.005% nicotinamide, 0.1% starch, 0.05% glucose, 0.9% NaCl, 0.23% leptospira base Ellinghausen-McCullaugh-Johnson-Harris (EMJH) (Difco), and 2% Noble agar (Difco). Immediately before being poured, this medium is supplemented with 0.0025% reduced NAD and 1% chicken serum. TM/SN base consists of 1% biosate peptone (BBL), 1% NaCl, 0.1% starch, 0.05% glucose, and 1.5% Noble agar (Difco) and is supplemented with 5% oleic albumin complex, 1% chicken serum, 0.0005% thiamine, and 0.0025% reduced NAD. A modified version of TM/SN which consists of brain heart infusion agar that is supplemented with 5% oleic albumin complex, 1% chicken serum, 0.0005% thiamine, and 0.0025% reduced NAD has proven to be as suitable as the original formula given above. Broth versions of HMM and TM/SN are prepared by omission of the agar.

Infectious Coryza

whereas A. paragallinarum will colonize and produce typical disease.

AGENT IDENTIFICATION Background Avibacterium paragallinarum is not the only growth factor­ dependent organism that can be isolated from chickens. Nonpathogenic avian hemophili have been recognized since the 1930s. The organisms have been the subject of several taxonomic changes - being initially named as Haemophilus avium (16) and then as Pasteurella volantium, Pasteurella avium, and Pasteurella species A (22). In a recent polyphasic study, these non-pathogenic organisms have been transferred to the genus Avibacterium as A. avium, A. volantium and Avibacterium species A (2). The final member of the new genus is A. (Pasteurella) gallinarum. A range of other hemophili, none of which are yet assigned to named species, have been isolated from birds other than chickens. The identification of these other hemophili will not be discussed further here.

Morphology Examination of a direct smear of the exudate by Gram stain can be useful for initial assessment of the microbial flora. The finding of gram-negative rods in direct smears, however, is not sufficient grounds for definitive diagnosis and must be followed by culture. After incubation of blood agar plates for 24—^48 hr, V factor­ dependent isolates of A. paragallinarum produce tiny dewdrop colonies up to 0.3 mm in diameter adjacent to the nurse culture. The colonies become smaller with increasing distance from the nurse culture. For the satellitic growth to be obvious, cultures must be examined within 24-48 hr. The V factor-independent A. paragallinarum isolates produce small colonies (1-2 mm) that do not show satellitic growth. Colonies of A. avium, A. volantium, Avibacterium species A show satellitic growth and are typically much bigger than V factor-dependent A. paragallinarum. Some isolates of A. volantium may produce a yellowish pigment. Growth Requirements Most avian hemophili have a requirement for V factor but not for X factor (6). The determination of the growth factor requirements of these organisms is not an easy process. The use of some brands of commercial growth factor discs on media such as brain-heart infusion agar or susceptibility test agar can result in a high percentage of cultures that falsely appear to be both X and V factor­ dependent. The combination of Oxoid growth factor discs (Unipath, Ltd., distr. Unipath, Ogdensburg, N.Y.) and TM/S (TM/SN without added NAD) has been shown to be suitable for growth factor testing (5). Some isolates of avian hemophili may have such lowered V factor requirements that the serum must be omitted from TM/S. Alternative tests such as the porphyrin test (18) for X factor testing or the use of purified hemin (X factor) and NAD (V factor) as supplements to otherwise complete media are possible but are generally too complex for diagnostic laboratories. V factor-independent isolates of A. paragallinarum have been recognized to date in the Republic of South Africa (21) and Mexico (15). Hence, diagnostic bacteriologists need to be aware of the possibility of such variants emerging in other areas. Typically, A. paragallinarum isolates fail to grow in air, although some strains will develop this ability on subculturing. A. avium, A. volantium and Avibacterium species A all grow vigorously in air.

Chicken Embryos Avian hemophili can be propagated in 5 to 7-day-old chicken embryos, commonly by inoculation via the yolk sac route. After overnight incubation, large numbers of hemophili are present in the yolk, which then can be harvested and preserved by freezing at -70 C (or lower) or by lyophilization. Chicken Inoculation Another efficient diagnostic procedure is to inoculate suspect exudate into the infraorbital sinus of two or three susceptible chickens (preferably 4 wk old or more). The appearance of the typical clinical signs of infectious coryza in 24-48 hr is diagnostic. On occasion, if the number of viable organisms is low, particularly in chronic cases, the incubation period may be delayed for up to 1 wk. In such cases, a second passage may be required to produce the typical rapid onset of clinical signs. If the exudate is heavily contaminated with extraneous bacteria, the chicken inoculation test can be more reliable than culture. In such instances, most of the extraneous bacteria will be cleared by the host defense mechanisms,

Physicochemical Properties The ability to reduce nitrate to nitrite and ferment glucose without the formation of gas is common to all of the avian hemophili. Oxidase activity, the presence of the enzyme alkaline phosphatase,

23

Pat J. Blackall

Kume serogroups, the use of absorbed antisera allows the recognition of serovars, with the nine currently recognized Kume serovars being A-l, A-2, A-3, A-4, B-l, C-l, C-2, C-3, and C-4 (4). The antisera, with the exception of the antisera for B-l and C-3, need to be absorbed to allow recognition of the Kume serogroups. The antisera to be absorbed are diluted 1 in 40 in PBS-B-G. The absorption is performed using antigens adjusted to 64 HA units and at five times die volume of the diluted serum. The adjusted antigen is centrifuged and the supernatant discarded. The pelleted antigen is resuspended in the diluted antiserum. The suspension is left at room temperature for 2 hr and then overnight at 4 C. The suspension is then centrifuged and the supernatant retained as the absorbed antiserum (still at 1 in 40 dilution). Depending on the antiserum, a succession of absorptions may need to be performed. Once the antigen absorption has been completed, the sera need to be absorbed with fixed erythrocytes, using 5 times the volume of fixed erythrocytes compared with the serum. The relevant volume of erythrocytes is centrifuged and the supernatant discarded. The pelleted erythrocytes are resuspended in the diluted antiserum. The suspension is left at room temperature for 2 hr and then overnight at 4C. The suspension is then centrifuged and the supernatant retained as the absorbed antiserum (still at 1 in 40 dilution). The Kume scheme has not been widely applied as it is technically demanding to perform.

and a failure to produce indole or hydrolyse urea or gelatin are also uniform characteristics. A. paragallinarum is catalase-negative, whereas all other members of the genus Avibacterium including the hemophilic species of A. avium, A. volantium and Avibacterium species A are catalase-positive (2). Carbohydrate fermentation patterns of the avian hemophili are generally determined using a phenol red broth (Difco) containing 1% NaCl, 0.0025% NADH, 1% chicken serum, and 1% carbohydrate. For routine identification, the use of this broth and a dense inoculation (i.e., the modified reference technique of Blackall (1)), is a most suitable approach for determining fermentation patterns. For larger studies, a replica plating technique is more suitable (1). Table 6.1 presents those properties that allow full identification of the avian hemophili. The properties of all members of the genus Avibacterium are presented. The failure of A. paragallinarum to ferment either galactose or trehalose clearly separates it from all other members of the genus, both hemophilic and non-hemophilic.

Serotyping Tests Two different, but interrelated, serotyping schemes for A. paragallinarum have been used mainly the Page (23) and the Kume (19) schemes. The most widely recognized and applied scheme is the Page scheme, which recognizes three serovars (A, B, and C) of A. paragallinarum . Although the original scheme was based on an agglutination test, the Page serovars are best recognized by a hemagglutination-inhibition (HI) test. The antigen for this HI test can be produced by either of two methods. In the first method, whole bacterial cells are harvested from an overnight broth and stored at 4 C in 1/100 of the initial broth volume for at least 3 days (3). Alternatively, the antigen can be produced by the technique originally used in the Kume scheme (19). In this method, cells of A. paragallinarum are harvested, washed in phosphate-buffered saline (PBS) (pH 7.0), resuspended in 0.5 M KSCN/0.425 M NaCl to a density equivalent to a MacFarland nephelometer tube number 5 (Difco), held at 4 C for 2 hr with agitation, and then sonicated. The antigen is washed three times in PBS and resuspended in PBS with 0.01% merthiolate to a density equivalent to a MacFarland nephelometer tube number 5. With either antigen type, the HI test should be performed with glutaraldehyde-fixed chicken erythrocytes. The erythrocytes are prepared by collecting fresh chicken blood into an equal volume of Alsever’s solution. The suspension is centrifuged and the erythrocytes are washed three times in 0.15 M NaCl. The erythrocytes are then suspended to 1% (v/v) in a glutaraldehyde-salts solution and held at 4 C for 30 min. The glutaraldehyde-salts solution is prepared by diluting 25% glutaraldehyde to 1% in a solution containing one volume of 0.15 M Na2HPO4 (pH 8.2), nine volumes of 0.15 M NaCl, and five volumes of distilled water. The fixed erythrocytes are collected by centrifugation, washed five times in 0.15 M NaCl and five times in distilled water and finally resuspended to 30% (v/v) in distilled water containing 0.01% merthiolate. This stock of fixed erythrocytes is held at 4 C, and a working dilution of 1% is prepared in PBS (pH 7.0) containing bovine serum albumin (0.1%) and gelatin (0.001%) (PBS-B-G). Reports that Page serovar B is not a true serovar (28) have been discounted, and Page serovar B has been conclusively shown to be serologically distinct (34). The Kume scheme is a hemagglutination-inhibition serotyping scheme using bacterial cells that have been treated with potassium thiocyanate and then sonicated, and glutaraldehyde-fixed chicken erythrocytes. The preparation of the antigen and the chicken erythrocytes for the Kume scheme has already been described in the section on the Page serotyping scheme. The nomenclature of the Kume scheme has been changed since the original publication. Under die altered terminology the three Kume serogroups are termed A, B, and C (4). This terminology emphasizes that the Kume serogroups correspond to the Page serovars. Within the

Molecular Identification A polymerase chain reaction (PCR) test that is specific for A. paragallinarum has been described (11). The test has been evaluated with a wide range of A. paragallinarum isolates and is specific and sensitive. The test, termed the HP-2 PCR, has been shown to be suitable for use on purified DNA extracts as well as on crude colony preparations obtained from isolation plates. The HP-2 PCR can be used directly on swabs of nasal mucus obtained from the squeezing of the nostril. The test can be inhibited in the presence of blood and swabs obtained via by slicing open the infra­ orbital sinus during necropsy are not optimal for use in the HP-2 PCR. Direct PCR examination of swabs has been shown to outperform culture in China (10, 12). The HP-2 PCR is particularly useful in regions where both NAD-independent A. paragallinarum and Omithobacterium rhinotracheale are present. Molecular typing methods such as restriction endonuclease analysis (7) and ribotyping (20) have proved useful in epidemiologic studies. A PCR-based typing method - ERIC-PCR - has also shown a capacity to sub-type isolates of A. paragallinarum (30).

Maintenance Although the initial isolation of A. paragallinarum from acute infectious coryza is not difficult, it is a fragile organism requiring special care for propagation and maintenance in the laboratory. Cultures can be maintained on blood agar plates by weekly passages. Cultures incubated for 24-48 hr at 37 C and then stored at 4 C in a candle jar will remain viable for up to 2 wk. Cultures can be preserved by the lyophilization or freezing (at -70 C or lower) of infected yolk (see the section on chicken embryos). Storage at -70 C of a heavy suspension in the commercial bead systems now commonly available is also possible. SEROLOGIC DETECTION IN THE HOST Although a range of serologic tests for the detection of antibodies to A. paragallinarum have been described (6), only hemagglutination-inhibition (HI) tests are in widespread use. The various HI tests differ in the methods used to prepare the antigen and the type of red blood cells used. For the purposes of this overview, the three main HI tests have been termed the simple, extracted, and treated HI tests. Although most of these HI tests were originally described as tube or macroplate tests, all can also be

24

Chapter 6 Infectious Coryza

(one volume of antigen with eight volumes of erythrocytes for 2 hr at 37 C with serum recovered by centrifugation and regarded as being a one in five dilution). The extracted HI test has been used to detect antibodies to Page serovars A, B, and C in vaccinated chickens (35). Vaccinated chickens with HI titers of 1:5 or greater in the HI tests based on the simple and extracted antigens have been found to be protected against subsequent challenge (29).

performed in microtiter trays using appropriate volumes. The simple HI test is based on whole bacterial cells of a Page serovar A organism (17). Cells are harvested and suspended in PBS containing 0.01% merthiolate (pH 7.0). Pretreatment of the sera may be necessary to eliminate nonspecific agglutinins. A 5% suspension of chicken erythrocytes is added to the sera (1:5 proportion) and the mixture is incubated for 2 hr at room temperature and 12 hr at 4 C (29). Two-tenths milliliter of antigen (containing 20 HA units/ml) is added to 0.2 ml of serially diluted serum (initial dilution of 1:5). After incubation at room temperature for 10 min, 0.4 ml of 0.5% chicken erythrocyte suspension containing 0.02% (v/v) gelatin is added. The HI titers are determined after incubation for 30-40 min at room temperature. This test has been performed mainly using antigen prepared from A. paragallinarum strain 221. The extracted HI test is based on potassium thiocyanate-extracted and sonicated cells of A. paragallinarum and glutaraldehyde-fixed chicken erythrocytes (29). The preparation of the antigen and the chicken erythrocytes has already been described in the section on the Page serotyping scheme. Although the methodology of the extracted HI test is as described for the simple HI test, the sera are pretreated with 10% glutaraldehyde-fixed erythrocytes to eliminate nonspecific hemagglutinins, and PBS containing 0.1% bovine serum albumin and 0.001% gelatin is used as the diluent. The extracted HI test has been used to detect antibodies in chickens receiving inactivated vaccines based on Page serovar C organisms (29). In chickens infected with a serovar C organism, the majority of the birds remain negative in this test (36). Note that, in this report, the erythrocytes were fixed in formalin instead of glutaraldehyde (36). The treated HI test is based on hyaluronidase-treated whole bacterial cells of A. paragallinarum and formaldehyde-fixed chicken erythrocytes (37). Whole bacterial cells that have been adjusted to 10 times the optical density of 0.4 at 540 nm are treated with hyaluronidase (50 units/ml) in phosphate buffer (pH 6.0) in a waterbath at 37 C for 2 hr. The treated antigen is then washed twice in PBS and then resuspended in the original volume of PBS. The test uses the same methodology as that described for the extracted HI test except that the erythrocytes are 1.0% formaldehyde-fixed chicken erythrocytes. The diluent used in this HI test is the same as that used in the extracted HI test. All sera are pretreated with 50% (v/v) formaldehyde-fixed chicken erythrocytes

DIFFERENTIATION FROM CLOSELY RELATED AGENTS

Infection with A. paragallinarum must be differentiated from other diseases, such as chronic respiratory disease, chronic fowl cholera, fowl pox, omithobacteriosis (due to O. rhinotracheale), swollen head syndrome (associated with avian pneumovirus) and hypovitaminosis A, which can produce similar clinical signs. Because A. paragallinarum often occurs in mixed infections, one should consider the possibility of other bacteria or viruses as complicating agents, particularly if mortality is high and the disease takes a prolonged course. The simple isolation of a satellitic, gram-negative organism is not sufficient to identify an isolate as A. paragallinarum. For laboratories with limited facilities, sufficient evidence for a diagnosis of infectious coryza would be the isolation of a catalase­ negative, gram-negative organism that exhibits satellitic growth and fails to grow in air, together with a history of a rapidly spreading acute coryza in a flock. The use of the PCR is recommended as the definitive test. Laboratories without access to PCR technology should determine growth factor requirements and the ability to ferment glucose, galactose, and trehalose. At this level, A. paragallinarum can be differentiated from the other members of the genus Avibacterium. For laboratories with extensive resources, a complete phenotypic identification based on the properties shown in Table 6.1 is possible. As V factor-independent A. paragallinarum have been isolated, diagnostic microbiologists must be aware of the fact that non-satellitic bacteria should be considered as suspect A. paragallinarum if they show the typical biochemical properties of the species and have been obtained from chickens showing upper respiratory disease.

Table 6.1. Distinguishing properties of the species within the genera A vibacteriumK Property

Avibacterium gallinarum

Avibacterium paragallinarum

Avibacterium volantium

Avibacterium avium

Avibacterium sp. A.

Catalase Symbiotic growth Growth in Air

+B +

V -

+ + +

+ + +

+ + +

Acid from L-arabinose D-galactose Lactose D-mannitol Maltose D-sorbitol Trehalose ONPG

+ V + + V

+ V +

+ V + + V + +

+ +

+ + V V + V

-

A All species are Gram-negative and non-motile. All species reduce nitrate, are oxidase positive and ferment glucose. Most isolates of Avibacterium paragallinarum require an enriched carbon dioxide (5-10%) atmosphere and most will show an improved growth in the presence of 5-10% chicken serum. Most isolates of Avibacterium gallinarum show an improved growth in an enriched carbon dioxide (5-10%) atmosphere.® + = positive (>90%), - = negative (>90%), V = variable reaction

25

Pat J. Blackall 17. Iritani, Y., G. Sugimori, and K. Katagiri. Serologic response to

ACKNOWLEDGEMENT

Haemophilus gallinarum in artifically infected and vaccinated chickens.

Avian Dis 21:1-8. 1977. 18. Kilian, M. A rapid method for the differentiation of Haemophilus strains. Acta. Pathol. Microbiol. Immunol. Scand. Sect. B 82:835-842. 1974. 19. Kume, K., A. Sawata, T. Nakai, and M. Matsumoto. Serological classification of Haemophilus paragallinarum with a hemagglutinin system J. Clin. Microbiol. 17:958-964. 1983. 20. Miflin, J. K., R. F. Homer, P. J. Blackall, X. Chen, G. C. Bishop, C. J. Morrow, T. Yamaguchi, and Y. Iritani. Phenotypic and molecular characterization of V-factor (NAD)-independent Haemophilus paragallinarum. Avian Dis 39:304-308. 1995. 21. Mouahid, Μ., M. Bisgaard, A. J. Morley, R. Mutters, and W. Mannheim. Occurrence of V-factor (NAD) independent strains of Haemophilus paragallinarum. Vet. Microbiol. 31:363-368. 1992. 22. Mutters, R., K. Piechulla, K.-H. Hinz, and W. Mannheim Pasteurella avium (Hinz and Kunjara 1977) comb. nov. and Pasteurella volantium sp. nov. Int. J. Syst. Bacteriol. 35:5-9. 1985. 23. Page, L. A. Haemophilus infections in chickens. 1. Characteristics of 12 Haemophilus isolates recovered from diseased chickens. Am. J. Vet. Res. 23:85-95. 1962. 24. Poemomo, S., Sutarma, M. Rafiee, and P. J. Blackall. Characterization of isolates of Haemophilus paragallinarum from Indonesia. Aust. Vet. J. 78:759-762. 2000. 25. Rimler, R. B. Studies of the pathogenic avian haemophili. Avian Dis 23:1006-1018. 1979. 26. Rimler, R B., E. B. Shotts Jr, J. Brown, and R. B. Davis. The effect of sodium chloride and NADH on the growth of six strains of Haemophilus species pathogenic to chickens. J. Gen. Microbiol. 98:349-354. 1977. 27. Sandoval, V. E., H. R. Terzolo, and P. J. Blackall. Complicated infectious coryza cases in Argentina. Avian Dis 38:672-678. 1994. 28. Sawata, A., K. Kume, and Y. Nakase. Biologic and serologic relationships between Page's and Sawata's serotypes of {THaemophilus paragallinarum}. Am. J. Vet. Res. 41:1901-1904. 1980. 29. Sawata, A., K. Kume, and Y. Nakase. Hemagglutinin of Haemophilus paragallinarum serotype 2 organisms: occurrence and immunologic properties of hemagglutinin. Am J. Vet. Res. 43:1311-1314. 1982. 30. Soriano, V. E., G. Tellez, B. M. Hargis, L. Newberry, C. SalgadoMiranda, and J. C. Vazquez. Typing of Haemophilus paragallinarum strains by using enterobacterial repetitive intergenic consensus-based polymerase chain reaction. Avian Dis 48:890-895. 2004. 31. Terzolo, H. R, F. A. Paolicchi, V. E. Sandoval, P. J. Blackall, T. Yamaguchi, and Y. Iritani. Characterization of isolates of Haemophilus paragallinarum from Argentina. Avian Dis 37:310-314. 1993. 32. Thitisak, W., O. Janviriyasopak, R S. Morris, S. Srihakim, and R. V. Kruedener. Causes of death found in an epidemiological study of native chickens in Thai villages. Proc. 5th. Inter. Sym Vet. Epidemiol. Economics pp. 200-202. 1988. 33. Tongaonkar, S., S. Deshmukh, and P. Blackall. Characterisation of Indian isolates of Haemophilus paragallinarum. In 51st Western Poultry Disease Conference/XXVII convention anual ANECA. Peurto Vallajarta, Mexico, p. 58. 2002 34. Yamaguchi, T., P. J. Blackall, S. Takigami, Y. Iritani, and Y. Hayashi. Pathogenicity and serovar-specific hemagglutinating antigens of Haemophilus paragallinarum serovar B strains. Avian Dis 34:964-968. 1990. 35. Yamaguchi, T., P. J. Blackall, S. Takigami, Y. Iritani, and Y. Hayashi. Immunogenicity of Haemophilus paragallinarum serovar B strains. Avian Dis 35:965-968. 1991. 36. Yamaguchi, T., Y. Iritani, and Y. Hayashi. Serological response of chickens either vaccinated or artificially infected with Haemophilus paragallinarum. Avian Dis 32:308-312. 1988. 37. Yamaguchi, T., Y. Iritani, and Y. Hayashi. Hemagglutinating activity and immunological properties of Haemophilus paragallinarum field isolates in Japan. Avian Dis 33:511-515. 1989.

The authors would like to acknowledge the contribution of Dick Yamamoto who has been the senior author or co-author for previous editions of this text REFERENCES

1. Blackall, P. J. An evaluation of methods for the detection of carbohydrate fermentation patterns in avian Haemophilus species. J. Microbiol. Methods 1.275-281. 1983. 2. Blackall, P. J., H. Christensen, T. Beckenham, L. L. Blackall, and M. Bisgaard. Reclassification of Pasteurella gallinarum, [Haemophilus] paragallinarum, Pasteurella avium and Pasteurella volantium as Avibacterium gallinarum gen. nov., comb, nov., Avibacterium paragallinarum comb, nov., Avibacterium avium comb. nov. and Avibacterium volantium comb. nov. International Journal of Systematic and Evolutionary Microbiology 55:353-362. 2005. 3. Blackall, P. J., L. E. Eaves, and G. Aus. Serotyping of Haemophilus paragallinarum by the Page scheme: comparison of the use of agglutination and hemagglutination-inhibition tests. Avian Dis 34:643-645. 1990. 4. Blackall, P. J., L. E. Eaves, and D. G. Rogers. Proposal of a new serovar and altered nomenclature for Haemophilus paragallinarum in the Kume hemagglutinin scheme. J. Clin. Microbiol. 28:1185-1187. 1990. 5. Blackall, P. J., and J. G. Farrah. An evaluation of commercial discs for the determination of the growth factor requirements of the avian haemophili. Vet Microbiol. 10:125-131. 1985. 6. Blackall, P. J., and M. Matsumoto. Infectious coryza. In: Diseases of Poultry, 11th ed. Y. M. Saif, H. J. Bames, J. R. Glisson, A. M. Fadly, L. R. McDougald, and D. A. Swayne, eds. Iowa State University Press, pp. 691703 7. Blackall, P. J., C. J. Morrow, A. Mclnnes, L. E. Eaves, and D. G. Rogers. Epidemiologic studies on infectious coryza outbreaks in northern New South Wales, Australia, using serotyping, biotyping, and chromosomal DNA restriction endonuclease analysis. Avian Dis 34:267-276. 1990. 8. Bland, Μ. P., A. A. Bickford, B. R. Charlton, G. C. Cooper, F. Sommer, and G. Cutler. Case Report: A severe infectious coryza infection in a multi­ age layer complex in central California. In 51st Western Poultry Disease Conference/XXVH convention anual ANECA. Peurto Vallajarta, Mexico, p. 56-57. 2002 9. Bragg, R. R., P. Jansen Van Rensburg, E. Van Heerden, and J. Albertyn. The testing and modification of a commercially available transport medium for the transportation of pure cultures of Haemophilus paragallinarum for serotyping. Onderstepoort J Vet Res 71:93-98. 2004. 10. Chen, X., Q. Chen, P. Zhang, W. Feng, and P. J. Blackall. Evaluation of a PCR test for the detection of Haemophilus paragallinarum in China. Avian Pathol. 27:296-300. 1998. 11. Chen, X., J. K. Miflin, P. Zhang, and P. J. Blackall. Development and application of DNA probes and PCR tests for Haemophilus paragallinarum. Avian Dis 40:398-407. 1996. 12. Chen, X., C. Song, Y. Gong, and P. J. Blackall. Further studies on the use of a polymerase chain reaction test for the diagnosis of infectious coryza Avian Pathol. 27:618-624. 1998. 13. Chen, X., P. Zhang, P. J. Blackall, and W. Feng. Characterization of Haemophilus paragallinarum isolates from China. Avian Dis 37:574-576. 1993. 14. Droual, R., A. A. Bickford, B. R. Charlton, G. L. Cooper, and S. E. Channing. Infectious coryza in meat chickens in the San Joaquin Valley of California. Avian Dis 34:1009-10016. 1990. 15. Garcia, A. J., E. Angulo, P. J. Blackall, and A. M. Ortiz. The presence of nicotinamide adenine dinucleotide-independent Haemophilus paragallinarum in Mexico. Avian Dis 48:425-429. 2004. 16. Hinz, K.-H., and C. Kunjara. Haemophilus avium, a new species from chickens. Int. J. Syst. Bacteriol. 27:324-329. 1977.

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7

Campylobacter INFECTIONS IN POULTRY Jaap A. Wagenaar and Wilma F. Jacobs-Reitsma

SUMMARY. Poultry may easily become colonized with Campylobacter jejuni and/or Campylobacter coli. These bacterial species do not cause clinical disease in poultry but contamination of the meat during laughter and processing is a well recognized source of foodbome illness in humans. Agent Identification. Campylobacter is isolated from poultry feces and cecal contents using selective agar media under microaerobic conditions. Subsequently, suspect isolates are confirmed to be Campylobacter and identified to the species level by using a limited number of biochemical tests or appropriate PCR tests. Serologic Detection in the Host. Campylobacter infections are not routinely diagnosed by serologic assays.

INTRODUCTION

CLINICAL DISEASE

The gut of poultry easily becomes colonized with the thermophilic Campylobacter species C. jejuni and C. coli. Other Campylobacter species are very rarely (< 1%) isolated from poultry. Also wild birds frequently are found to carry Campylobacter spp., with a strong association of C. lari present in seagulls. Campylobacters are commensal organisms without causing any clinical disease in poultry, though there is an immune response and the presence of the organism is detected by the host. The significance of Campylobacter in poultry is for reasons of human public health and not for reasons of clinical poultry diseases. Vertical transmission of the infections does not occur, and eggs are not contaminated. Contamination of meat products is the main issue and consequently C. jejuni and C. coli are of importance only in broilers and other meat producing avian species. Many broiler flocks are colonized at the age of slaughter because poultry can be infected with low numbers of Campylobacter and Campylobacter is generally present in the environment with reservoirs in both domesticated and wild warm-blooded animals (18, 32). The fact that the organism colonizes in high concentrations results in shedding of high levels of Campylobacter and intense contamination of the environment. Cleaning and disinfection of poultry houses and the surroundings of the houses between production cycles is difficult. When the houses are stocked with a new flock of day-old chickens, the infection cycle can easily start again. When Campylobacter-^QsitiNQ birds are processed in the slaughterhouse, the meat may easily become contaminated with bacteria from the gut contents. Campylobacter will only multiply under specific atmospheric conditions with reduced oxygen tension and within a temperature range of 30 - 45 C. These conditions are not present during processing, transport and storage of the meat. In contrast with e.g. Salmonella, the Campylobacter concentration will not increase along the food chain in case of troubles with maintaining chilled conditions. At the consumer level, Campylobacter will easily die off during proper heating of the meat but the risk for humans is mainly in cross contamination from the raw meat to ready-to-eat products like salads. Campylobacter is a highly infectious organism so even ingesting low doses is associated with a relatively high probability of human illness. In addition to foodbome infections, humans may become ill from direct contact with infected animals like poultry, but also pets like (young) cats and dogs. Petting zoos are identified as a risk factor especially when less attention is paid to personal hygiene. Campylobacter is one of the most important causes of bacterial gastro-enteritis in humans (30). Campylobacteriosis in humans is characterized by diarrhea and abdominal cramps. Complications that may occur are reactive arthritis and the Guillain-Barre syndrome (10, 25).

Sporadic cases of vibrionic hepatitis in poultry have been described, supposedly caused by Campylobacter. However, there is only the suggestion of a causative role of Campylobacter without any proof (7). C. jejuni may cause illness in ostriches but the economic loss due to Campylobacter infections are assumed to be very limited (28). Both C. jejuni and C. coli have a high prevalence in poultry, but both species generally are considered as normal gut inhabitants. Co­ infections with multiple strains frequently occur. There is a strong seasonality in the prevalence in broiler flocks with higher isolation rates in summer (18). Control programs in poultry to reduce the number of infected flocks are implemented to prevent human illness from contaminated poultry meat. Epidemiology of Campylobacter infections in broiler flocks is still not completely elucidated. Recently a systematic review based on UK data and comprising 159 research papers was published on risk factors for introduction of Campylobacter into broiler flocks (1). Partial depopulation (thinning) and multiple poultry houses on a farm were identified as contributing factors associated with increased risk, and hygiene barrier, parent company and certain seasons of rearing were associated with decreased risk.

Diagnostics Testing poultry for presence of Campylobacter is performed for particular research objectives, for more general monitoring reasons (observing trends), or to identify the Campylobacter status of the individual flock. The latter may be used to decide on scheduled slaughter of negative flocks followed by positive flocks. Monitoring programs Monitoring programs are implemented to identify trends in Campylobacter infections and to evaluate the feasibility of control programs. It offers the possibility to link the poultry data to the human Campylobacter data in order to assess the contribution of poultry to the human burden of illness. A good example is the obligatory monitoring of Campylobacter in broilers in the EU as prescribed by Zoonoses Directive 2003/99/EC. According to this directive the monitoring of broiler flocks started on January 1st, 2005 with the European Food Safety Authority (EFSA) as the responsible agency for compilation and reporting of data collected by the EU member states. At this moment (February 2006) the EU is unique in the world for this monitoring program. (http ://www.efsa. eu.int/science/monitoring zoonoses/reports/catind ex en.html). Individual flocks In some European countries (Denmark, Iceland, Norway) poultry flocks are tested for Campylobacter prior to slaughter, and negative and positive flocks are slaughtered separately. The meat from positive flocks is treated (freezing or heat treatment) to reduce the Campylobacter concentration, whereas the meat from negative

27

Jnp A Wagenaar and Wilma F. Jacobs-Reitsma

flocks is sold as fresh meat. As consumers are exposed to reduced levels of Campylobacter compared to the situation without separation, this approach aims to reduce the burden of illness. The effect of this approach is strongly dependent on the reliability (in particular sensitivity) of the diagnostic procedure, including collection and transport of samples, and performance of the detection assay.

fecal samples is not performed routinely, enrichment media will not be discussed in this chapter. Several commercial enzyme immunoassays are available for the detection of Campylobacter in human stool samples but these tests are not validated for routine use in poultry fecal samples.

Selective media for isolation Many media currently are in use for the bacteriological culture of Campylobacter spp. and the majority also are commercially available. A recommendation for a specific medium cannot be given. Experience in laboratories is an important factor in the choice of the medium. There are differences in medium preferences in different places in the world. Many European countries use mCCDA whereas in the US Campy-cefex and CVA are more commonly used (see list below). A detailed description of the developments in Campylobacter detection and the variety of existing media is given by Corry et al. (8, 9). Campylobacterselective media can be divided into two main groups: blood­ containing media and charcoal-containing media. Both blood components and charcoal remove toxic oxygen derivatives. The selectivity of the media is determined by the antimicrobials used. Cefalosporins (generally cefoperazone) are used, sometimes in combination with other antibiotics (e.g. vancomycin, trimethoprim). Cycloheximide (actidione) or amphotericin B are used to inhibit yeasts and molds (17). All media allow the growth of C. jejuni and C. coli but the main difference between the media is the degree of inhibition of contaminating flora depending on the combination of antimicrobials used. There is no medium available that allows growth of C. jejuni and inhibits C. coli or vice versa. To some extent, other Campylobacter species (e.g. C. lari, C. upsaliensis, C. helveticus, C. fetus and C. hyointestinalis) will also be able to grow on some of these media, especially at the less selective temperature of 37 C. Examples of selective blood-containing solid media are: Preston agar (4), Skirrow agar (24), Campy-cefex (29), CVA (22). Examples of selective charcoal-based solid media are:mCCDA (modified Charcoal Cefoperazone Deoxycholate Agar)(ll), slightly modified version of the originally described CCDA (5), Karmali agar or CSM (Charcoal-Selective Medium) (14), CAT agar (Cefoperazone, Amphotericin, Teicoplanin agar), facilitating growth of C. upsaliensis (2).

SAMPLE COLLECTION

Isolation of Campylobacter from feces and contents of the ceca is described here as these samples are commonly used for the diagnosis of a Campylobacter infection in poultry. Carcass samples can be analyzed but this needs a specific approach including selective enrichment. Isolation of Campylobacter from carcass and meat samples is described in detail elsewhere (12,13). Collection of samples A Campylobacter infection rapidly spreads within a flock (31), and close to 100% of the animals become colonized, shedding >106 cfu Campylobacter per gram feces for a prolonged period of time (at least till slaughter age of broilers). At a within-flock prevalence of >40%, 10 samples will be sufficient to detect the Campylobacter infection at a 95% confidence level. In case of scheduled slaughter it is most important to know the actual Campylobacter status of the flock just prior to slaughter. Consequently, the samples should be taken as close to slaughter as possible: at the farm or even at the slaughterhouse. In the latter case, reliable results can be obtained by taking intact ceca for examination. Living birds can be sampled by taking fecal/cecal droppings or cloacal swabs. For reliable detection of Campylobacter by culture, freshly voided feces should be collected. All samples must be prevented from drying out before culture. At the slaughterhouse, ceca can be cut with sterile scissors from the remaining part of the intestines and submitted intact to the laboratory in a plastic bag or Petri-dish.

Transport of samples Campylobacter easily dies off during transport of samples and precautions have to be taken to prevent the samples from dehydration, atmospheric oxygen, sunlight and elevated temperature. When swabs are used, a transport medium (like Amies, Cary Blair or Stuart) must be used. These transport swabs are available commercially. When only small amounts of fecal/cecal samples can be collected and transport swabs are not available, shipment of the specimen in transport media is recommended. Several transport media have been described: Cary-Blair, modified Cary-Blair, modified Stuart medium, Campythioglycolate medium, alkaline peptone water and semisolid motility test medium. Good recovery results have been reported using Cary-Blair (15, 23). No specific recommendation on the temperature for transportation can be made, but it is clear that freezing and high temperatures reduce viability. In general high temperatures (>20 C), low temperatures ( 1:20 in larger birds are frequently seen in cases of recent infection. A negative result does not guarantee that a bird is free of infection as the sensitivity of the test is rather low. The direct complement fixation (DCF) test detects avian IgG but not IgM. An advantage is that there is a readily available microprocedure. Disadvantages are: 1) test antigen commercially unavailable, 2) the test cannot be used for testing sera from avian species whose immunoglobulins do not fix complement, such as small psittacine birds, 3) it is only relatively sensitive, 4) it cannot be used to differentiate between IgG and IgM antibodies, making it necessary to test paired sera, and 5) the technique is fairly laborious when there is a large number of samples to be tested. The modified DCF test is more sensitive but has the same disadvantages as the DCF test (5,21,23,47). The indirect IF test detects all immunoglobulin isotypes and is, as is the MIF test, widely used to detect Chlamydia trachomatis, Chlamydophila pneumoniae and Chlamydophila psittaci antibodies in human sera using selected strains of these bacteria (40,55,17). The MIF test appears to be more sensitive than the complement fixation tests. Some years ago a large number of commercially available ELISAs were evaluated for demonstrating Chlamydophila psittaci antibodies in birds. All of these ELISAs were highly sensitive but showed low specificity, as they were mainly based on the use of whole chlamydial organisms, LPS, or chlamydial outer membrane fractions of LPS and lipoglycoprotein nature (44,7,18,30,47). When using these sources and antigen, false-positives due to the presence of antibodies crossreactive to the chlamydial LPS or hsp60 cannot be ruled out. More recently, peptide-based ELISA systems, or ELISAs using recombinant LPS, have become commercially available for the specific detection of Chlamydia trachomatis, Chlamydophila pneumoniae and Chlamydophila abortus antibodies (Medac, Savyon, Labsystems), (28,34,26,46. These tests performed as well as the MIF assay, but are less time-consuming, less expensive, and easier to perform. In the future this principle might also be useful in

the serodiagnosis of Chlamydophila psittaci infections. At present, an ELISA using recombinant MOMP of Chlamydophila psittaci has already been described (52; Verminnen et al., 2005, unpublished results) for testing avian sera. The test is not species-specific as the recombinant MOMP, produced by transiently tranfected COS7 cells (51), comprises all four variable domains as well as conserved domains and thus detects antibodies against all members of the genus Chlamydophila and Chlamydia (53). However, this is not a problem when testing avian sera as birds can only become infected by Chlamydophila psittaci. Unfortunately, Chlamydophila psittacispecific ELISAs are not yet available.

DIFFERENTIATION FROM CLOSELY RELATED AGENTS

The gross lesions caused by chlamydiae may resemble those caused by Pasteurella, mycoplasmas, and coliforms when a systemic infection occurs. Pasteurella- and coliform-caused diseases can readily be eliminated from consideration, as the fibrinous exudate from uncomplicated chlamydial infection will be free of bacteria that can be cultured on standard bacteriologic media. Confirmation of chlamydial infections can be obtained by isolation and identification of viable chlamydiae from infected tissues or by demonstration of chlamydiae by specific immunohistochemical techniques.

Table 16.1. Modified Giminez technique or Pierce-Van Der Kamp (PVK) stain.

Constituent

Amount

Solution 1:

Distilled H2O Phenol

450.0 ml 5.0 ml

Add to:

Basic fuchsin 95% Ethanol

2-5 g 50.0 ml

Incubate (tightly capped) at 37 C for 48 hr. Filter and store at room temperature. Solution 2:

Na2HPO4 NaH.POAO Distilled H2O (pH 7.5)

Solution 3:

Solution 1 Solution 2

11.65 g 2.47 g Sufficient quantity 1 liter 20.0 ml 25.0 ml

Let stand 10 min, filter, and use.

Solution 4:

0.5% Citric acid (aqueous)

Solution 5:

Fast green Distilled H2O Glacial acetic acid

Solution 6:

Solution 5 Distilled H2O

Chlamydiosis

0.2 g 100.0 ml 0.2 ml 20.0 ml 50.0 ml

71

Arthur A. Andersen and Daisy Vanrompay

Table 16.2 Primer sequences for the nested PCR/EIA Oligonucleotide

bp

Forward outer

21

CCT GTA GGG AAC CCA GCT GAA

Reverse outer

22

GGT TGA GCA ATG CGG ATA GTA T

Fluorescein—forward inner

17

GCA GGA TAC TAC GGA GA

Biotin—reverse inner

18

GGA ACT CAG CTC CTA AAG

Sequence (5’- 3’)

Table 16.3. Genotype specific primers and probes and competitors Oligonucleotide

Sequence (5’-3’)

Positiona

Specificity

CpPsSSfor

TTATTAAGAGCTATTGGTGGATGCC

1 822

Cp. psittaci

CpPsSSrev

AACGTATAATGGTAGATGATTAATCTACCG

1 972

CpPsGASfor

GGTTTTCAGCTGCAAGCTCAA

488

CpPsGASprobe

CTACCGATCTTCCAACGCAACTTCCTAACG

512

CpPsGAScompetitorB

CTACCGATCTTCCAATGCAACTTCCTAACGb

512

CpPsGASrev

CCACAACACCTTGGGTAATGC

565

CpPsGBSfor

AATAGGGTTTTCAGCTACCAACTCAA

483

CpPsGBSprobe

TCTACCGATCTTCCAATGCAACTTCCTAACGTA

511

CpPsGBScompetitorA

TCTACCGATCTTCCAACGCAACTTCCTAACGTA

511

CpPsGBScompetitotE+E/B

TCTACCGAGCTTCCAATGCAACTTCCTAACGTA

511

CpPsGBSrev

CCACAACACCTTGGGTAATGC

565

CpPsGCSfor

GCATCGCTCAACCTAAATTGG

929

CpPsGCSprobe

TCTGCTGTTATGAACTTGACCACATGGAACC

952

CpPsGCSrev

ATTGTGGCTTCCCCTAAAAGG

1 009

CpPsGDSfor

AACCACTTGGAACCCAACACTTT

969

CpPsGDSprobe

AGGAAAGGCCACAACTGTCGACGG

993

CpPsGDSrev

CGAAGCAAGTTGTAAGAAGTCAGAGTAA

1 062

CpPsGESfor

CCAAGCCTTCTAGGATCAAGGA

982

CpPsGESprobe

TACTTTGCCCAATAATGGTGGTAAGGATGTTCTATC

1 005

CpPsGEScompetitorA+B

TGCTTTGCCCAATAATAGTGGTAAGGATGTTCTATC

1005

CpPsGEScompetitorE/B

TGCTTTGCCCAATAATGCTGGTAAGGATGTTCTATC

1 005

CpPsGESrev

CGAAGCAATTTGCAAGACATCA

1 062

CpPsGFSfor

GCAACTTTTGATGCTGACTCTATCC

904

CpPsGFSprobe

CATCGCTCAACCTAAATTAGCCGCTGC

930

CpPsGFSrev

GTTCCATGTGGTCAAGTTCAAAAC

981

CpPsGE/BSfor

CCAAGCCTTCTAGGATCAACCA

982

CpPsGE/BSprobe

TGCTTTGCCCAATAATGCTGc

1 005

CpPsGE/BScompetitorA+B

TGCTTTGCCCAATAATAGTG

1 005

CpPsGE/BScompetitorE

TACTTTGCCCAATAATGGTG

1 005

CpPsGE/BSrev

TGCAAGACATCAGATAGAACATCCTT

1 052

72

Genotype A

. Genotype B

Genotype C

Genotype D

Genotype E

Genotype F

Genotype E/B

Chapter 16

ML-INR03-B

ML-INF02-F

872 bp

< 70

Chlamydiosis

> 638

ompA VD1

CD2

VD2

920

655

CD3

VD3

CD4

941 VD4

1068 bp

CD5

—►

Figure 16.1: Location of the outer and inner primers in the ompA gene. Numbering according to ompA sequences in Genbank

15. Everett, K.D.E., RM. Bush, and A.A. Andersen. Emended description of the order Chlamydiales, proposal of Parachlamydiaceae fam nov. and Simkaniaceae fam. nov., each containing one monotypic genus, revised taxonomy of the family Chlamydiaceae, including a new genus and five new species, and standards for the identification of organisms. Inti. J. Systematic Bacteriol. 49:415-440. 1999. 16. Everett, K.D.E., L.J. Hornung, and A.A. Andersen. Rapid detection of the Chlamydiaceae and other families in the Order Chlamydiales'. Three PCR Tests. J. Clin. Microbiol. 37(3):575-580. 1999. 17. Fernandez, F., J. Gutierrez, J. Mendoza, J. Linares, and M.J. Soto. A new microimmunofluorescence test for the detection of Chlamydia pneumoniae specific antibodies. J. Basic Microbiol. 44(4):275-9. 2004. 18. Fudge, A.M. Blocking antibody ELISA testing of pet birds: Implications for chronic infections. Semen. Avian Exotic Pet Med. 2:167— 170. 1993. 19. Geens, T., A. Dewitte, N. Boon, and D. Vanrompay. Development of a Chlamydophila psittaci species—specific and genotype—specific real-time PCR. Vet. Res. 2005, in press. 20. Geens, T., A. Desplanques, M. Van Loock, B.M. Bonner, E.F. Kaleta, S. Magnino, A.A Andersen, K.D. Everett, and D. Vanrompay. Sequencing of the Chlamydophila psittaci ompA gene reveals a new genotype, E/B, and the need for a rapid discriminatory genotyping method. J. Clin. Microbiol. 43(5):2456—61. 2005. 21. Grimes, J.E., D.N. Phalen, and F. Arizmendi. Chlamydia latex agglutination antigen and protocol improvement and psittacine bird antichlamydial immunoglobulin reactivity. Avian Dis. 37(3):817-24. 1993. 22. Grimes, J.E., and F. Arizmendi. Usefulness and limitations of three serologic methods for diagnosing or excluding chlamydiosis in birds. J. Am. Vet. Med. Assoc. 209:747-750. 1996. 23. Grimes, J.E., T.N. Tully, Jr., F. Arizmendi, and D.N. Phalen. Elementary body agglutination for rapidly demonstrating chlamydial agglutinins in avian serum with emphasis on testing cockatiels. Avian Dis. 38:822-831. 1994. 24. Grimes, J.E. Evaluation and interpretation of serological responses in psittacine bird chlamydiosis and suggested complementary diagnostic procedures. J. Avian Med. Surg. 10:75—83. 1996. 25 Hewinson, R.G., P.C. Griffiths, B.J. Bevan, S.E. Kirwan, M.E. Field, M.J. Woodward, and M. Dawson. Detection of Chlamydia psittaci DNA in avian clinical samples by polymerase chain reaction. Vet. Microbiol. 54(2): 155-66. 1997. 26. Hoymans, V.Y., J.M. Bosmans, L. Van Renterghem, R. Mak, D. Ursi, F. Wuyts, C.J. Vrints, and M. Ieven. Importance of methodology in determination of Chlamydia pneumoniae seropositivity in healthy subjects and in patients with coronary atherosclerosis. J. Clin. Microbiol. 41(9)4049-53. 2003. 27. Kaltenbock, Β., N. Schmeer, and R. Schneider. Evidence for numerous ompl alleles of porcine Chlamydia trachomatis and novel chlamydial species obtained by PCR. J. Clin. Microbiol. 35(7): 1835-41. 1997.

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Arthur A. Andersen and Daisy Vanrompay

28. Kaltenboeck, B., D. Heard, F.J. DeGraves, and N. Schmeer. Use of synthetic antigens improves detection by enzyme—linked immunosorbent assay of antibodies against abortigenic Chlamydia psittaci in ruminants. J. Clin. Microbiol. 35(9):2293-8. 1997b. 29. Kutyavin, I.V., I.A. Afonina, A. Mills, V.V. Gom, E.A. Lukhtanov, E.S. Belousov, MJ. Singer, D.K. Walburger, S.G. Lokhov, A.A. Gall, R. Dempcy, MW. Reed, R.B. Meyer, and J. Hedgpeth. 3'-minor groove binder-DNA probes increase sequence specificity at PCR extension temperatures. Nucleic Acids Res. 28:655-661. 2000. 30. Ley, D.H., K. Flammer, and P. Cowen. Performance characteristics of diagnostic tests for avian chlamydiosis. J. Assoc. Avian Vet. 7:203-207. 1993. 31. Lindqvist, R., B. Norling, and S.T. Lambertz. A rapid sample preparation method for PCR detection of food pathogens based on buoyant density centrifugation. Lett. Appl. Microbiol. 24:306—310. 1997. 32. Madico, G., T.C. Quinn, J. Boman, and C. A. Gaydos. Touchdown enzyme time release—PCR for detection and identification of Chlamydia trachomatis, C. pneumoniae, and C. psittaci using the 16S and 16S—23 S spacer rRNA genes. J. Clin. Microbiol. 38(3): 1085-93. 2000. 33. Messmer, T. O., S. K. Skelton, J. F. Moroney, H. Daugharty, and B. S. Fields. Application of a nested, multiplex PCR to psittacosis outbreaks. J. Clin. Microbiol. 1997; 35:2043-2046. 34. Morre, S.A., C. Munk, K. Persson, S. Kruger-Kjaer, R. van Dijk, C.J. Meijer, and A. J. van Den Brule. Comparison of three commercially available peptide-based immunoglobulin G (IgG) and IgA assays to microimmunofluorescence assay for detection of Chlamydia trachomatis antibodies. J. Clin. Microbiol. 40(2):584-7. 2002. 35. Moore, F.M, and ML. Petrak. Chlamydia immunoreactivity in birds with psittacosis: Localization of chlamydiae by the peroxidase­ antiperoxidase method. Avian Dis. 29:1036-1042. 1985. 36. Paul, I.D. The growth of Chlamydia in McCoy cells treated with emetine. Med. Lab. Sci. 39:15-32. 1982. 37. Ridstrom, P., R. Knutsson, P. Wolffs, M Dahlenborg, and C. Lofstrom. Pre—PCR processing of samples. In: PCR detection of microbial pathogens. K. Sachse, and J. Frey, eds. Humana Press, Totowa, NJ. pp. 31-50. 2003. 38. Sachse, K. and H. Hotzel. Detection and differentiation of Chlamydiae by nested PCR. In: PCR detection of microbial pathogens. K. Sachse and J. Frey, eds. Humana Press, Totowa, NJ. pp. 123-136. 2003. 39. Sachse, K, H. Hotzel, P. Slickers, T. Ellinger, and R. Ehricht. DNA microarray-based detection and identification of Chlamydia and Chlamydophila spp. Mol. Cell Probes 19(1):41—50. 2005. (Epub 2004 Nov Π) 40. Salinas, J., MR. Caro, and F. Cuello. Comparison of different serological methods for the determination of antibodies to Chlamydia psittaci in pigeon sera. Zentralbl. Veterinarmed. B. 40(4):239—44. 1993. 41. Sayada, C., A.A. Andersen, C. Storey, A. Milon, F. Eb, N. Hashimoto, K. Hirai, J. Elion, and E. Denamur. Usefulness of ompl restriction mapping for avian Chlamydia psittaci isolate differentiation. Res. Microbiol. 146:155-165. 1995. 42. Smith, K.A., K.K. Bradley, MG. Stobierski, and L.A. Tengelsen. Compendium of measures to control Chlamydophila psittaci (formerly Chlamydia psittaci) infection among humans (psittacosis) and pet birds, 2005. J. Am. Vet. Med. Assoc. 226(4):532-39. 2005.

43. Spencer, W.N., and F.W.A. Johnson. Simple transport medium for the isolation of Chlamydia psittaci from clinical material. Vet. Rec. 113:535— 536. 1983. 44. Sting, R., and H.M. Hafez. Purification of Chlamydia psittaci antigen by chromatografie on polymyxin B agarose for use in die enzyme-linked immunosorbent assay (ELISA). Zentralbl. Bakteriol. 277: 436—445. 1992. 45. Tappe, J.P., A.A. Andersen, and N.F. Cheville. Respiratory and pericardial lesions in turkeys infected with avian or mammalian strains of Chlamydia psittaci. Vet. Pathol. 26:386-395. 1989. 46. Tiran, A., K. Tiesenhausen, E. Karpf, J. Orfila, G. Koch, H.J. Gruber, O. Tsybrovskyy, and B. Tiran. Association of antibodies to chlamydial lipopolysaccharide with the endovascular presence of Chlamydophila pneumoniae in carotid artery disease. Atherosclerosis. 173(1):47—54. 2004. 47. Tully, T.N.,Jr., S.M. Shane, J.E. Grimes, R.P. Poston, and MT. Kearney. Comparison of procedures to detect Chlamydia psittaci antibodies in cockatiels (Nymphicus hollandicus). Avian Dis. 40(2): 266—71. 1996. 48. Van Loock, Μ, K. Verminnen, T.O. Messmer, G. Volckaert, B.M Goddeeris and D. Vanrompay. Use of a nested PCR-enzyme immunoassay with an internal control to detect Chlamydophila psittaci in turkeys. 2005, BMC Infect. Dis. 5:76, 2005. 49. Vanrompay, D., R. Ducatelle, and F. Haesebrouck. Diagnosis of avian chlamydiosis: Specificity of the modified Gimenez staining on smears and comparison of the sensitivity of isolation in eggs and three different cell cultures. J. Vet. Med. Ser. B 39:105-112. 1992. 50. Vanrompay, D., A. Van Nerom, R. Ducatelle, and F. Haesebrouck. Evaluation of five immunoassays for detection of Chlamydia psittaci in cloacal and conjunctival specimens from turkeys. J. Clin. Microbiol. 32:1470-1474. 1994. 51. Vanrompay, D., E. Cox, J. Mast, B. Goddeeris, and G. Volckaert. Highlevel expression of Chlamydia psittaci major outer membrane protein in COS cells and in skeletal muscles of turkeys. Infect. Immun. 66(11): 5494500. 1998. 52. Vanrompay, D., E. Cox, P. Kaiser, S. Lawson, M Van Loock, G. Volckaert, and B. Goddeeris. Protection of turkeys against Chlamydophila psittaci challenge by parenteral and mucosal inoculations and the effect of turkey interferon-gamma on genetic immunization. Immunology. 103(1): 106-12. 2001. 53. Vanrompay, D., T. Geens, A. Desplanques, T.Q. Hoang, L. De Vos, M Van Loock, E. Huyck, C. Mirry, and E. Cox. Immunoblotting, ELISA and culture evidence for Chlamydiaceae in sows on 258 Belgian farms. Vet. Microbiol. 99(1):59—66. 2004. 54. Wilson, P.A., J. Phipps, D. Samuel, andN.A. Saunders. Development of a simplified polymerase chain reaction—enzyme immunoassay for the detection of Chlamydia pneumoniae. J. Applied Bacteriol. 80:431—438. 1996. 55. Wong, K.H, Skelton S.K., and H. Daugharty. Utility of complement fixation and microimmunofluorescence assays for detecting serologic responses in patients with clinically diagnosed psittacosis. J. Clin. Microbiol. 32(10):2417-21. 1994. 56. Yoshida, Η., Y. Kishi, S. Shiga, and T. Hagiwara. Differentiation of Chlamydia species by combined use of polymerase chain reaction and restriction endonuclease analysis. Microbiol. Immunol. 42(5):411—4. 1998.

17 ORNITHOBACTERIOSIS Richard P. Chin and Bruce R. Charlton

SUMMARY. Omithobacteriosis is a respiratory disease of avian species caused by the bacterium Ornithobacterium rhinotracheale. Gross lesions in infected birds include a fibrinoheterophilc pneumonia, airsacculitis, and pericarditis. Ornithobacterium rhinotracheale is a pleomorphic gram-negative rod-shaped bacterium that grows on blood agar. Seventeen serotypes (A through Q) are currently recognized, though serotype A is the major one isolated. The bacterium has been isolated from numerous avian species, primarily infecting chickens and turkeys. Agent Identification. Isolation and identification of the bacterium are required to make a diagnosis. Serologic Detection in the Host. Enzyme-linked immunosorbent assays and serum plate agglutination tests have been developed to detect antibodies.

INTRODUCTION

airsacculitis, pericarditis, peritonitis, and a mild tracheitis could occur.

Ornithobacterium rhinotracheale is associated with respiratory disease in avian species, primarily turkeys and chickens, but the bacterium also has been isolated from chukars, rooks, and a partridge, pheasant, and pigeon (2,4,10). The first reported isolation of O. rhinotracheale was made from turkeys in Germany in 1981 (6). The bacterium has since been isolated from birds throughout the world. Prior to being named in 1994, O. rhinotracheale had been identified as Pasteurella-\&Q organism, Kingella-tiks bacterium, Taxon 28, and pleomorphic gram-negative rod bacterium. Identification of O. rhinotracheale by biochemical test kits can be challenging. Current commercially available identification systems do not have O. rhinotracheale listed in their database and may misidentify it as Gallibacterium anatis biovar haemolytica (formerly Pasteurella haemolytica), Weeksella spp., or Flavobacterium meningosepticum, depending on the test system and variable reactions of this bacterium.

SAMPLE COLLECTION

The primary site for isolating O. rhinotracheale is the respiratory tract. Taking tracheal cultures from live birds should be done using a sterile swab. Care must be taken to prevent contamination as large numbers of other bacteria can easily overgrow the pinpoint colonies of O. rhinotracheale. Samples can be taken at necropsy using sterile swabs (cotton or dacron) from the trachea, infraorbital sinuses, lungs, and air sacs. Reports have been made of isolation of O. rhinotracheale from joints, heart, brain, and fiver, and in chronic disease, from the thoracic vertebrae and tendon sheaths. Isolates can be stored at -70 C. PREFERRED CULTURE MEDIA AND SUBSTRATES For primary isolation, O. rhinotracheale gjtows readily on 5% sheep blood agar. It grows aerobically, microaerobically, and anaerobically. The best growth occurs in air enriched with 7.5%10% CO2 at 37 C, although growth occurs from 30 to 42 C. Ornithobacterium rhinotracheale does not grow on MacConkey agar. About 90% of the isolates appear to be resistant to gentamicin and polymyxin B (11). Hence, 5pg/ml each of gentamicin and polymyxin B can be added to blood agar media for selective isolation of O. rhinotracheale. However, blood agar plates without antibiotics should always be used in tandem to prevent missing the 10% antibiotic-susceptible isolates.

CLINICAL DISEASE Whether or not O. rhinotracheale is a primary pathogen is still uncertain. In many cases, in both chickens and turkeys, infection with O. rhinotracheale is secondary to other respiratory disease agents (9). In chickens, infections have been reported in 3-to-6-wk-old birds, and in layers 20-50 wk of age (5,6,8,9,12). Mild respiratory signs occur from about 3 to 4 wk of age, mortality mildly increases, and airsacculitis condemnation rates at the slaughter plant are higher. In broiler breeders, the disease primarily occurs between 20 and 50 wk of age, during peak egg production (5). Mortality is slightly increased, feed intake is decreased, and mild respiratory signs occur. There may be egg production decrease, eggshell may be of poor quality, and egg size may decrease. In turkeys, infections have been detected as early as 2 wk of age, but severe lesions are seen in older birds (>14wk of age) and breeders (3,5,11). Infection with O. rhinotracheale at 2 wk of age causes respiratory signs and nasal discharge, followed by facial edema and swelling of the infraorbital sinuses (9). Birds will appear depressed with ruffled feathers, and display a decrease in feed and water intake. Subsequently, mortality increases. In older birds, a sudden increase in mortality may be the only sign. Slight depression and gasping, marked dyspnea, and expectoration of blood-stained mucus can be seen just prior to death. In breeder flocks, egg production can decrease slightly (2-5%) (3). Mortality rates usually range between 2% and 11% in chickens and turkeys. Clinical cases reveal unilateral and bilateral lung consolidation with fibrinous exudate on the pleura in both chickens and turkeys (3,11). These lesions are similar to those seen in fowl cholera in turkeys, but are less severe. In addition, a fibrinoheterophilic

AGENT IDENTIFICATION

Colony and Cell Morphology On blood agar, pinpoint colonies, less than 1 mm, can be detected at 24 hr of incubation. At 48 hr, colonies will be approximately 1-2 mm in diameter, gray, circular, and convex with an entire edge. Some isolates from chickens have a reddish glow. Plates should be held for 48 h to look for the small colonies typical of O. rhinotracheale as they can easily be overgrown by other bacteria such as Escherchia coli. Ornithobacterium rhinotracheale cultures can have a distinct smell similar to butyric acid. Ornithobacterium rhinotracheale is a gram-negative, highly pleomorphic, nonmotile, nonsporulating bacterium. It appears primarily as short, plump rods 0.2-0.9 pm x 1-3 pm, and less frequently as long filamentous rods or club-shaped rods (10). Biochemical Characteristics Biochemical tests for O. rhinotracheale are inconsistent. Listed in Table 17.1 are some of the more consistent reactions.

75

Richard P. Chin and Bruce R. Charlton

Table 17.1. Some phenotypic characteristics of Ornithobacterium

DIFFERENTIATION FROM CLOSELY RELATED AGENTS

rhinotracheale.

Test

Reaction

Blood agar MacConkey agar Gram stain Catalase Oxidase Triple sugar iron agar β-Galactosidase (ONPG) Indole

Growth No growth Pleomorphic gram-neg. rod Negative Positive No change Positive Negative

Ornithobacterium rhinotracheale infection should be differentiated from other diseases with pneumonia and airsacculitis, such as fowl cholera, colibacillosis, mycoplasmosis, and chlamydophylosis. This is dependent upon isolation and identification of O. rhinotracheale from infected tissues. REFERENCES

1. Back, A, D. Halvorson, G. Rajashekara, K. V. Nagaraja. Development of a serum plate agglutination test to detect antibodies to Ornithobacterium rhinotracheale. J. Vet. Diagn. Invest. 10:84-86. 1998 2. Charlton, B. R., S. E. Channing-Santiago, A. A. Bickford, C. J. Cardona, R. P. Chin, G. L. Cooper, R Droual, J. S. Jeffrey, C. U. Meteyer, H. L. Shivaprasad, and R. L. Walker. Preliminary characterization of a pleomorphic gram-negative rod associated with avian respiratory disease. J. Vet. Diagn. Invest. 5:47-51. 1993. 3. Chin, R. P., P. C. M van Empel, and Η. M Hafez. Ornithobacterium rhinotracheale infection. In: Diseases of poultry, 11th ed. Y. M Saif, H. J. Bames, J. R. Glisson, A. M Fadly, L. R. McDougald, and D. E. Swayne, eds. Iowa State University Press, Ames, Iowa. pp. 683-690. 2003.

The API-20NE identification strip (bioMerieux-Vitek, Hazelwood, Mo.) is routinely used, although O. rhinotracheale is not included in their database, and there are various biocodes. The most common biocodes are 0220004 (61%) and 0020004 (38.5%) (11). If the isolate is positive for arginine dihydrolase (0.5%) biocodes of 0320004 and 0120004 are produced. The API ZYM system (bioMerieux-Vitek, Hazelwood, Mo.) is useful for determining enzymatic activities. In this system O. rhinotracheale consistently produces five negative reactions (lipase, β-glucuronidase, βglucosidase, α-mannosidase, and α-fucosidase). Another rapid test system, the RapID NF Plus, (Remel/Atlanta, Norcross, Ga.) also appears to be suitable for identification of O. rhinotracheale (7). Field isolates generated 5 unique biocodes: 472264 (41.8%), 476264 (31.8%), 676264 (18.2%), 672264 (7.3%) and 472044 (0.9%). Carbohydrate fermentation is extremely variable. Analysis of cellular fatty acids can aid in identifying O. rhinotracheale. Predominant fatty acids detected using the Microbial Identification System (Microbial ID, Newark, Del.) are 15:0 iso, 16:0, 15:0 iso 3OH, 17:0 iso, 16:0 3OH, 17:0 iso 3OH, and unknown peaks with equivalent chain lengths of 13.566 and 16.580 (2). Seventeen serotypes of O. rhinotracheale are reported, serotype A through Q. These have been differentiated using the agar gel immunodiffusion test (8), (P. van Empel, pers. comm.). Some geographical differences exist in serotypes found throughout the world. Most isolates found in Europe and the United States are serotype A. After serotype A, types B, D, and E are the predominate serotypes found in Europe. Ninety-five percent of the strains isolated from chickens are serotype A.

4. De Rosa, M., R. Droual, R. P. Chin, H. L. Shivaprasad, and R. L. Walker. Ornithobacterium rhinotracheale infection in turkey breeders. Avian Dis. 40:865-874. 1996. 5. Hafez, Η. M Current status on the role of Ornithobacterium rhinotracheale (ORT) in respiratory disease complexes in poultry. Arch. Geflugelk. 60:208-211. 1996. 6. Hinz, K.-H., and Η. M Hafez. The early history of Ornithobacterium rhinotracheale (ORT). Arch. Geflugelk 61:95-96. 1997. 7. Post, K. W., S. C. Murphy, J. B. Boyette, and P. M Resseguie. Evaluation of a commercial system for the identification of Ornithobacterium rhinotracheale. J. Vet. Diagn. Invest. 11:97-99. 1999. 8. Travers, A. F. Concomitant Ornithobacterium rhinotracheale and Newcastle disease infection in broilers in South Africa. Avian Dis. 40:488490. 1996. 9. van Beek, P.N.G.M, P. C. M van Empel, G. Van Den Bosch, P. K. Storm, J. H. Bongers, and J. H. du Preez. Ademhalingsproblemen, groeivertraging en gewrichtsontsteking bij kalkoenen en vleeskuikens door een Pasteurella-achtige bacterie: Ornithobacterium rhinotracheale or “Taxon 28.” Tijdschr. Diergeneeskd 110:99-101. 1994. 10. Vandamme P., P. Segers, M. Vancanneyt, K. van Hove, R. Mutters, J. Hommez, F. Dewhirst, B. Paster, K. Kersters, E. Falsen, L. A Devriese, M Bisgaard, K.-H. Hinz, and W. Mannheim. Ornithobacterium rhinotracheale gen. nov., sp. nov., isolated from the avian respiratory tract. Int. J. Syst. Bacteriol. 44:24-37. 1994. 11. van Empel, P. C. M, and Η. M Hafez. Ornithobacterium rhinotracheale: a review. Avian Pathol. 28: 217-227. 1999. 12. van Empel, P., H. van den Bosch, D. Goovaarts, and P. Storm. Experimental infection in turkeys and chickens with Ornithobacterium rhinotracheale. Avian Dis. 40:858-864.1996.

SEROLOGIC DETECTION IN THE HOST Enzyme-linked immunosorbent assays (ELISAs) have been developed and appear adequate for detecting antibodies to most serotypes of O. rhinotracheale (5,8,11). Commercial ELISA kits are available in Europe and appear to detect antibodies for serotypes A through M. A serum plate agglutination test has also been developed to detect antibodies to O. rhinotracheale (1).

76

18 MYCOSES AND MYCOTOXICOSES R. D. Wyatt

SUMMARY. Mold-related disease in avian species can be divided into two broad categories, namely mycoses and mycotoxicoses. Mycoses are typically defined as infection of tissue by a particular mold species. In general terms, Aspergillus, Dactylaria, and Microsporum are those mold species most apt to be responsible for mycoses in avian species. Additionally, Candida, a genus of polymorphic yeasts (exhibiting yeast cells, hyphae and pseudohyphae) can also infect the upper gastrointestinal tract. All of these agents should be considered economically important to domestic poultry, however, Aspergillus is by far the most common. Mycotoxins are a broad and diverse group of toxic metabolites produced by toxigenic molds when these molds are permitted to grow on certain feedstuffs. Consumption of such moldy feedstuffs by avian species may result in clinical signs of disease, dependent upon the specific mycotoxin that is contaminating the ingested feedstuff. The three most important mold genera that are known to have species that posses the capability to produce mycotoxins are Fusarium, Aspergillus, and Penicillium. Trichothecenes, a rather large group of mycotoxins with similar chemical structures, are produced primarily by species of Fusarium. Aflatoxin, perhaps the most toxic of all mycotoxins, is the most important mycotoxin produced by Aspergillus. Ochratoxin is also an economically important mycotoxin and is produced by species of Penicillium and Aspergillus. Isolation and identification of molds responsible for mycoses can be accomplished in infected tissues by employing basic microbiological techniques. Additionally, histological examination of infected tissue and examination of the etiological agent and tissue response can also confirm the identity of die pathological agent. Isolation and identification of specific mycotoxins requires extraction of the contaminated feedstuff followed by chemical analysis of the extract. Additionally, flock history, management practices and a working knowledge of specific clinical signs of various mycotoxicoses is essential to identify the specific mycotoxin involved in a disease outbreak.

ASPERGILLOSIS INTRODUCTION dramatically. Consequently, the signs of mycotoxicoses will vary depending on the specific mycotoxin(s) involved in the disease. Signs of disease are also influenced by various interacting factors such as the presence of other diseases, nutritional status of affected birds, environmental conditions, and husbandry practices (11, 13, 17, 34, 35, 37, 39). Hence, a rapid and accurate diagnosis of specific mycotoxicoses is often very difficult.

The term “fungus” is a general term used to describe any member of a very broad taxonomic classification of eukaryotic microorganisms (protists) including lichens, mushrooms, yeasts, rusts, smuts, molds, and numerous other organisms. These organisms have no chlorophyll, are aerobic, and have a very rigid cell wall containing chitin. Molds are a subset of fungi that degrade organic material in the environment. They are ubiquitous, saprophytic, parasitic, and their spores are resistant to many environmental conditions, especially desiccation. When growing in culture, most molds will first produce single filaments known as hyphae and as the culture matures it will appear filamentous and assume a "wooly” appearance. Mature cultures are frequently pigmented due to the production of pigments by the mycelium or pigments associated with spores. Pathogenic molds typically reproduce by formation of spores or conidia; whereas, Candida albicans reproduces by budding. A relatively small percentage of molds in die environment are causative of disease of poultry. Nevertheless, based upon routine exposure of poultry to molds, the probability for disease outbreaks to occur in commercial poultry is significant. Infections of the upper respiratory tract by Aspergillus fumigatus and other Aspergilli and infections of the upper gastrointestinal tract of poultry by Candida albicans are welldocumented (7,40). Other mold species are merely environmental inhabitants with soil being their natural ecological niche. These molds are typically associated with degradation of living or dead organic material. Those degrading living organic material are often plant pathogens; whereas, those degrading dead organic material are best known for their capability to produce secondary metabolites such as antibiotics (e.g. Penicillium chrysogenum). Some molds possess the genetic capability to produce highly toxic secondary metabolites during their growth on organic material. These toxic metabolites are referred to as mycotoxins. Ingestion of mycotoxins by poultry in grain or finished feed will result in disease referred to as mycotoxicosis. Mycotoxicoses in poultry are quite diverse with each being caused by a specific mycotoxin and each toxicosis characterized by a primary target organ system, target organ, or target tissue. The toxicity of various mycotoxins to poultry differ

CLINICAL DISEASE Aspergillosis can be caused by any member of the genus Aspergillus, however, Aspergillus fumigatus is the most common etiologic agent for this disease in poultry (7, 18, 22). The lungs and air sacs are the typical tissues involved, however, infection of the eye, sinuses, and meninges can occur. When responsible for mycoses, Aspergilli are generally considered opportunistic pathogens. Immunosuppression, respiratory tract irritation, antibiotic therapy and die presence of other infective agents can predispose birds to aspergillosis (2). In young broilers, poults, and quail, and a variety of wild birds (7, 12, 22) high morbidity and mortality are commonly observed. Improperly fumigated hatching eggs and improperly disinfected incubators and hatchers can result in high exposure rates to the spores of Aspergilli resulting in infection of chicks with the disease progressing rapidly. Consequently, the term “brooder pneumonia” is often used to describe acute aspergillosis in young birds. Impairment of respiratory function followed by lethargy and anorexia can lead to death in young birds within 24-72 hr. Adult turkeys are also susceptible to aspergillosis. Although the signs of the disease are similar to those of young turkeys, chickens and quail, the disease is generally of a longer duration. Coughing, sneezing, and general labored breathing are commonly observed. Upon necropsy, the internal lesions can be described as focal caseous nodules on the air sacs, imbedded within the lungs and on the mucosal surface of the trachea. Occasionally, greenish mold growth can be visualized on the air sacs. This is typically associated with the chronic form of the disease. When this occurs, the lesions associated with other internal organs may appear yellowish in color. 77

R. D. Wyatt

SAMPLE COLLECTION

AGENT IDENTIFICATION

Upon necropsy, areas of suspected infected tissue should be carefully identified, aseptically excised from surrounding tissue and immediately transferred to a sterile vessel. Retrieval of suspect tissue from both the periphery and center of a lesion is recommended. Retrieved tissue should be subjected to both direct microscopic examination and microbiological culturing. Care should be taken in maintaining aseptic conditions since Aspergilli are common in the laboratoiy microflora. When samples are being prepared for microscopic examination, immediate fixing of the specimen is imperative. Also, culturing procedures should be initiated without delay following removal of the infected tissue.

Presumptive identification of Aspergillus is based upon microscopic morphology and growth on various microbiological media. Aspergilli typically exhibit hyaline (glassy appearance), septate, and branched hyphae. The color of the conidiophore can be colorless, brown or green depending upon the species (7). The conidiophore with be rather long (> 300 microns) in the case of A. niger, and short (80% CPE, the cells are disrupted by freezing and thawing, the cell debris removed by centrifugation, and the supemates, containing virus antigens, concentrated by precipitation with ammonium sulphate overnight or by extraction with arcton. A control adenovirus antiserum and antigen is required and common lines of precipitation in agar, between the control and test antigen confirms presence of adenovirus. Growth of group 3 (EDS) virus in cell culture is accompanied by production of a hemagglutinin which agglutinates avian erythrocytes but not mammalian erythrocytes. Therefore, while fluorescence or other antigen detecting techniques can be applied to detection of EDS virus, hemagglutination presents an easier alternative. This is carried out by removing some supernatant from the EDS virus-infected culture, following development of CPE, and addition of an equal volume of chicken or duck erythrocytes (0.8% suspension in PBS). Presence of EDS hemagglutinin results in hemagglutination of the erythrocytes. Hemagglutination-inhibition (HI) (see below) using EDS virus-specific antiserum confirms the isolate as EDS virus. If immunofluorescence is to be used for detection of EDS virus antigen in infected cultures or tissues, antisera with specificity for group I adenoviruses cannot be used, because of lack of cross reactivity. Specific EDS antisera are required for this purpose. EDS specific antisera have been used for direct immunostaining of tissue sections to detect EDS antigens. Using the avidin-biotin-peroxidase technique, EDS antigen was detected in tissue sections of infundibulum for 3-5 days post infection and in large amounts in the pouch shell gland from about 8 days post inoculation, until as long as affected eggs were produced (19).

Nucleic acid detection Presence of adenovirus in tissues or lesions from diseased birds is often first suspected by demonstration of intranuclear inclusion bodies in histopathology studies, although adenoviruses are not the only agents which give rise to intranuclear inclusions following infection. Other viruses e.g. herpesviruses, circoviruses or polyoma viruses may give rise to similar inclusions. In situ hybridization (ISH) has been used for detection of adenovirus DNA in tissues in these circumstances and can be useful in species other than turkeys or chickens where cell cultures are not available for virus isolation, or where formalin-fixed tissues are the only specimen submitted to the laboratory for diagnosis. Adenovirus ISH probes (36-40 bases) based on a highly conserved region of the penton of FAdVIO have been described and used in ISH (12). This region was chosen since it is highly conserved and therefore provides maximum cross reactivity between adenovirus species. The procedure requires the sections to be de-paraffinized by treatment with xylene, rehydrated by passage through graded alcohols, digested with pepsin, before hybridization to the digoxigen labeled probe. An anti-digoxygenin antibody conjugated to alkaline phosphatase was then applied, and the chromogen, nitroblue tetrazolium (NBT) added. Foci of dye reduction to blue­ black formazan pigment indicated presence of adenovirus nucleic acid, which duplicated the pattern of intranuclear inclusions seen in parallel H&E stained sections. ISH was successfully used to demonstrate hepatic, renal and intestinal adenovirus infection in parrots and to exclude herpesvirus and circovirus from the diagnosis. The same probes were used ‘in an investigation of inclusion body hepatitis and pancreatitis in broiler chickens, and positive adenovirus ISH results were confirmed by thin section electron microscopy and virus isolation (10).

Morphology On development of CPE, negative stain electron microscopy is used to confirm presence of adenovirus. Simple techniques for diagnostic use of the electron microscope are described in Chapter 34 on enteric viruses. Examination of negatively stained material by electron microscopy is fast and also allows both positive identification of the virus and recognition of other unsuspected viruses or bacteria. Adenoviruses are normally easily recognized by their morphology. They are seen as isosahedral particles (i.e. particles with 12 vertices and 20 triangular faces) measuring 65-75 nm in diameter. Degenerating particles, however, can develop a circular appearance. Adeno-associated parvoviruses are frequently seen, along with the adenoviruses. These are spherical particles measuring approximately 22 + 3 nm in diameter. If desired, sensitivity can be improved by use of immunoelectron microscopy, in which adenovirus antiserum or convalescent serum is mixed with the specimen at an appropriate dilution to facilitate clumping of particles, making viewing easier. Thin-section electron microscopy has also proved useful for demonstration of adenoviruses in cell cultures or to confirm presence of adenovirus in tissues or organs from birds in which intranuclear inclusions are seen with high frequency in histopathology studies. Cell cultures showing CPE are scraped from the plastic surface, fixed for 2 hr at 4 C in glutaraldehyde, post fixed in osmium tetroxide for 2 hr at room temperature, then dehydrated in alcohol and embedded in Araldite. Ultrathin sections of infected cells stained with uranyl acetate and lead citrate show large numbers of characteristic particles, 65-75 nm in diameter, often in crystalline arrays, along with associated inclusion material in the nucleus (5). Small pieces of tissue shown by histopathology to contain large numbers of inclusions can also be treated in this way, and often show similar large concentrations of virus particles. This can be useful in investigations in species other than chicken and turkey where virus isolation in cell culture is more difficult, or not 86

Chapter 19 Adenovirus

an option, but is not indicative of a causal relationship to the condition.

virus stock is mixed with buffer at pH 3 and pH 7. After incubating for 1 hour at 37 C, the virus suspensions are immediately titrated. Adenoviruses should be stable to pH 3 treatment, and show no substantial reduction in infectivity. Adenoviruses in cell-culture medium, without serum, are generally inactivated by heating for 30 min at 60 C. At 50 C for 1 hr, 1 M MgCl2 enhances the thermal inactivation, and this is the basis of the test for stability in the presence of divalent cations.

Subtyping of Isolates The hexon protein of the group 1 avian adenoviruses contains a highly conserved region which gives rise to antibodies which are ‘group specific’ i.e. the antibodies produced following infection with one group 1 adenovirus also react with other group 1 viruses in tests such as gel diffusion and immunofluorescence. Other antibodies are produced following infection, which react only with the infecting virus and closely related viruses. These type specific antibodies neutralize virus infectivity and are used to separate group 1 viruses into serotypes, using the serum neutralization test (see below). In most cases, establishment an isolate as an adenovirus may be sufficient for diagnosis. Further characterization, to establish relatedness to standard strains, requires more detailed laboratory testing. The group 1 adenoviruses are subdivided into 5 species (A to E; Table 19.1), based largely on molecular criteria, in particular restriction endonuclease analysis (REA) and sequencing data. Viruses within each species are further subdivided into serotypes largely on the result of cross-neutralization tests, although this can also be done by PCR. REA of fowl adenovirus serotypes into 5 distinct groups was first described by Zsak and Kisary, (21) and has proved effective in clarifying relationships between serotypes and in some cases between strains showing differences in virulence e.g. strains associated with IBH (9). Polymerase chain reaction (PCR) primers, based on conserved regions of the hexon gene, are available which will amplify group 1 adenoviruses belonging to all 12 reference serotypes (15). Treatment of the PCR product with 4 endonucleases produces restriction patterns which are specific for each reference strain. Therefore, the combined use of PCR with REA can be used for detection and typing of all group 1 adenoviruses (15,11). Details of the primers and methodology are given by Meulemans et al., (15), and Hess (11). PCR’s are also available for specific detection of EDS virus (17) and for amplification of DNA from all 3 groups of avian adenoviruses (21). Subdivision into serotypes can also be done by serum neutralization test, which requires substantial cell culture expertise and access to reference antisera prepared against each prototype strain. Prototype antisera (usually prepared in rabbits) are titrated against 100 mean tissue culture infective doses (TdD50) of virus and then diluted to contain 20 neutralizing units (e.g., a serum with a neutralizing antibody titer of 1:2048 is diluted to 1:100). The strains are described in Table 19-1. For typing of isolates by serum neutralization, the virus isolate is diluted to 1/10 and 1/1000 in cell culture medium, mixed with 1 volume of each prototype antiserum containing 20 neutralizing units, and incubated for 2 hr at 37 C. Duplicate monolayers of chick embryo liver or kidney cells, growing in 96-well, flat bottomed microtitre plates or in 24 well cell culture plates, are inoculated with each virus-serum mixture and the plates are incubated at 37 C and observed for CPE over a period of 6 days. Controls, consisting of virus alone, and antiserum alone are also inoculated. Neutralization of virus CPE indicates the serotype of the isolate. Chick embryo liver cells are preferable to chick kidney cells because the endpoints are more distinct. If desired, pools of antisera can be used instead of individual antiserum.

SEROLOGIC DETECTION IN THE HOST

Group specific antibodies Because antibodies to avian adenoviruses are widespread in poultry flocks, serological methods are of little significance in diagnosis, except for the screening of SPF flocks. For this purpose, ELISA is probably the test of choice. Indirect ELISAs have been described and have been shown to be broadly group specific, detecting antibodies to all group 1 avian adenoviruses (7). Antigen for ELISA was grown by infecting monolayers of chicken kidney cells with each avian adenovirus serotype. After 4 days, the cells were scraped from the plastic, disrupted by sonication in distilled water and extracted with fluorocarbon. The antigen preparation was added to ELISA cuvettes at a predetermined dilution and allowed to absorb overnight. After washing, test antisera were added, the cuvettes were again washed and an anti-chicken serum conjugated with horseradish peroxidase was added. Following incubation and washing positive reactions were identified by addition of the enzyme substrate, at a previously standardized concentration (7). A similar indirect ELISA for detection of group 3 (EDS) virus antibodies has also been described (5). In this test the EDS antigen was grown in chick embryo liver cells, and an anti-chicken serum conjugated with peroxidase was used with ortho phenylene diamine as the substrate. The test was shown to be equal in sensitivity to an EDS serum neutralization test and an indirect fluorescent antibody test which used EDS virus infected chick embryo liver cells, fixed on 12-well multispot slides. A group specific, microtitre based immunofluorescence test capable of screening large numbers of sera for group 1 adenovirus antibodies, has also been described (4). In this test, chick embryo liver cells, grown as monolayers in the wells of 96-well, flat bottomed microtitre plates for 48 hr, were infected with FAdVl virus, for 24 hr, and the plates were fixed by addition of a diluted hypochlorite solution. After washing in PBS, test sera were applied and the plates were incubated at 37 C for 30 min then washed and an anti-chicken serum conjugated with FITC added. After incubation and washing, the wells were examined for fluorescent nuclei using an inverted fluorescence microscope. Double-immunodiffusion or gel diffusion has been commonly used for detecting avian adenovirus group specific antibodies. It is fast, cheap, and simple to use but lacks sensitivity, so birds undergoing a primary infection by natural routes may not respond with detectable precipitating antibodies. However, if three antigens are pooled (e.g. FAdVl, FAdV5 and FAdV9) sensitivity can be markedly improved (8). Antigen for gel diffusion test may be prepared by inoculating embryonating chicken eggs by the allantoic route with 0.1 ml of an egg-adapted strain of group 1 adenovirus, such as FAdVl. After 72 hr incubation at 37 C, the eggs are chilled, and the chorioallantoic membranes harvested. One ml PBS is added for each membrane and the membranes are homogenized then centrifuged at 1500 x g for 20 min to remove the debris. The pooled supemate is stored in small volumes at -80 C as gel diffusion antigen.

Physicochemical properties The more important physical and chemical properties may be useful if an isolate is recovered, which cannot be neutralized using the prototype antisera. Tests for determining the presence of essential lipid, nucleic acid type, and thermal stability may be useful and are described in Chapter 45 on virus identification and classification. Other characterization criteria may include morphology, site of replication, resistance to pH 3, and stability in the presence of divalent cations. To test for resistance to pH 3, the

Type specific (neutralizing) antibodies Neutralizing antibodies to group 1 adenoviruses may occur in chickens, turkeys, quail, pigeons, and other avian species. The neutralization test is similar to that described above for typing of 87

Brian M. Adair and J. Brian McFerran

titration in cell culture should be performed with new isolates. Ten­ fold dilutions of the virus isolate are prepared and inoculated into duplicate chick embryo liver or kidney cell cultures. The highest dilution showing CPE is harvested by freezing at -80 C and thawing; the process is repeated a total of 3 times. Electron microscope examination should be used to confirm presence of only one agent. While the procedure is not foolproof, it gives some confidence that most other agents have been removed. Adenoassociated parvoviruses have been found in adenovirus stock pools and have also been isolated from healthy birds. They require presence of adenovirus for their replication, and there is no evidence to suggest an involvement in disease. Molecular tests (PCR) are available for their diagnosis.

isolates. Two-fold dilutions of the test sera (e.g. 1/16 to 1/2048) are added (50ul volumes) to duplicate wells of flat-bottomed microtitre plates. Two hundred TCID50 ,of the appropriate adenovirus serotype in a 50 μΐ volume, are then added to each well. Following incubation for 1 hour at 37 C, to allow neutralization to take place, 50μ1 of chick embryo liver or kidney cell suspension is added to each well. Virus controls containing 100, 10, 1.0, and 0.1 TCID50 of virus are also included, and the plates are incubated and examined for development of viral CPE. Neutralizing endpoints are read when the virus controls indicate that 100 TCID5o of virus is present in the test. The same test can be used for EDS virus to confirm results of HI tests. In this case the endpoints can be read on the basis of CPE or by removing the supemate from each well to a parallel microtitre plate, adding 50μ1 of chicken erythrocytes and observing for presence of hemagglutination, as described above.

REFERENCES

1. Adair, B.M Studies on the development of avian adenoviruses in cell cultures. Avian Pathol. 7:541-550. 1978 2. Adair, B.M, Curran, W.L. and McFerran, J.B. Ultrastructural studies of the replication of fowl adenoviruses in primary cell cultures. Avian Pathol. 8:133-144. 1979. 3. Adair, B.M, McFerran, J.B., Connor, T.J., McNulty, MS. and McKillop, E.R. Biological and physical properties of a virus (strain 127) associated with the egg drop syndrome 1976. Avian Pathol. 8: 249-264. 1979. 4. Adair, B.M, McFerran, J.B. and Calvert, V.M Development of a microtitre fluorescent antibody test for serological detection of adenovirus infection in birds. Avian Pathol. 9: 291-300. 1980. 5. Adair, B.M., Todd, D., McFerran, J.B. and McKillop, E.R. Comparative serological studies with egg drop syndrome virps. Avian Pathol. 15: 677685. 1986. 6. Benko, M, B.Harrach, and W.C. Russell. 2000. Family Adenoviridae. In: Virus Taxonomy. Seventh Report of the International Committee on Taxonomy of Viruses. MH.V. van Regenmortel, C.M Fauquet, D.H.L. Bishop, E.B. Carstens, MK. Estes, S.M Lemon, J. Maniloff, MA. Mayo, D.J. McGeoch, C.R. Pringle, R.B. Wickner (eds), Academic Press, New York, San Diego. Pp. 227-238. 2000. 7. Calnek, B.W., Shek, W.R., Menendez, N.A. and Stiube, P. Serological cross-reactivity of avian adenovirus serotypes in an enzyme-linked immunosorbent assay. Avian Dis. 26: 897-906 1982. 8. Cowen, B. S. A triple antigen for the detection of type 1 avian adenovirus precipitin. Avian Dis. 31:351-354. 1987. 9. Emy, K. M, D. A Barr, and K. J. Fahey. Molecular characterization of highly virulent fowl adenoviruses associated with outbreaks of inclusion body hepatitis. Avian Pathol. 20:597-606. 1991. 10. Goodwin, MA, Latimer, K.S., Resurreccion, R.S., Miller, P.G. and Campagnoli, R.P. DNA in situ hybridization for the rapid diagnosis of massive necrotizing avian adenovirus hepatitis and pancreatitis in chicks. Avian Dis. 40:828-831. 1996. 11. Hess, M Detection and differentiation of avian adenoviruses: a review. Avian Pathol. 29: 195-206. 2000. 12. Latimer, K.S., Niagro, F.D., Williams, O.C., Ramis, A., Goodwin, MA, Ritchie, B.W. and Campagnoli, R.P. Avian Dis. 41: 773-782. 1997 13. McFerran, J.B. and Smyth, J.A Avian Adenoviruses. Rev. Sci. Off. Int. Epiz., 19: 589-601. 2000. 14. Mazaheri, A, Prusas, C., Voss, M and Hess, M Some strains of serotype 4 fowl adenoviruses cause inclusion body hepatitis and hydropericardium syndrome in chickens. Avian Pathol. 27: 269-276. 1998. 15. Meulemans, G., Boschmans, M., van den Berg, T.P. and Decaesstecker, M Polymerase chain reaction combined with restriction enzyme analysis for detection and differentiation of fowl adenoviruses. Avian Pathol. 30: 655660. 2001. 16. Nakamura, K., Mase, M, Yamaguchi, S., Shibahara, T. and Yuasa, N. Pathologic study of specific pathogen free chicks and hens inoculated with adenovirus isolated from hydropericardium syndrome. Avian Dis. 43: 414423. 1999. 17. Raue, R. and Hess, M Hexon based PCR’s combined with restriction enzyme analysis for rapid detection and differentiation of fowl adenoviruses and egg drop syndrome virus. J. Virol. Meth. 73: 211-217. 18. Saifuddin, M and Wilks, C.R. Development of an enzyme linked immunosorbent assay to detect and quantify adenovirus in chicken tissues. Avian Dis. 34: 239-245. 1990.

HI test for group 3 (EDS) virus antibodies EDS virus growth in cell cultures or eggs is accompanied by production of a hemagglutinating antigen, which agglutinates chicken and duck erythrocytes. Antibody to the virus inhibits the hemagglutination reaction and this is used as the basis of the HI test for EDS. The test can be used for detecting infection with EDS virus and is not applicable to any other avian adenovirus. Birds do not usually develop antibody until after they have had EDS. Flocks that develop EDS through reactivation of latent virus usually do so before they are 35 wk old. Birds of all ages are susceptible to the virus, and if the EDS virus is introduced (e.g., through drinking contaminated water), infection can occur at any age. For screening purposes, 1/10 dilutions of test sera in PBS are added to duplicate wells of V-well microtitre plates. An equal volume of EDS antigen, diluted to contain 4 hemagglutinating (HA) units, is added to one well, and 1 volume PBS to the duplicate (control) well. A titration of a positive serum (2-fold dilutions in PBS) should also be included. The plates are shaken and left at room temperature for 15 min, 1 volume of an 0.8% suspension of chicken erythrocytes is added to each well. The plates are again shaken and left at room temperature until the erythrocytes have settled (about 45 min), and then viewed by holding at an angle of 45 degrees and allowing the settled erythrocytes to stream. No hemagglutination should occur in the serum control well, no inhibition should occur with negative serum, and the positive serum should be at its correct titer. Finally, a back-titration carried out on the diluted antigen used in the test should indicate that 4 HA units are present. Antigen for HI tests may be prepared by inoculating 10-day-old embryonated duck eggs allantoically with 0.2 ml of a 1/100 dilution of EDS virus. The allantoic fluid is harvested 4-5 days after inoculation, clarified by low-speed centrifugation, and stored at -80 C in small volumes. Alternatively chick embryo liver or chick embryo kidney monolayers may be inoculated with 1000 TCID50 of EDS virus, and incubated until EDS virus CPE is seen in 10% of the cells. The flasks of cells are then frozen at -80 C and thawed 3 times and the cell debris removed by centrifugation. The supernatant containing the hemagglutinin is distributed in small volumes and stored frozen at -80 C. If inactivation of the antigen is required, this is done by heating for 1 hr at 65 C in a water bath. This is preferred to formalin treatment.

DIFFERENTIATION FROM CLOSELY RELATED AGENTS Differentiation of Isolates. Isolation attempts in cell cultures will often result in the isolation of other agents, particularly reoviruses. In addition to the different CPE in cells, these viruses produce intracytoplasmic inclusions. Mycoplasma may also be recovered from specimens in cell culture and may be suspected due to their reovirus-like CPE and sensitivity to chloroform. To eliminate the possibility of contamination with other agents, limit dilution 88

Chapter 19

Adenovirus

20. Xie, Z., Fadi, A.A., Girshick, T. and Khan, MI. Detection of avun adenovirus by polymerase chain reaction. Avian Dis. 43: 98-105. 1999. 21. Zsak, L. and Kisary, J. Grouping of fowl adenoviruses based upon the restriction patterns of DNA generated by Bam Hl and Hind HI. InterviroL 22: 110-114.

19. Smyth, J. A., M A. Platten, and J. B. McFerran. A study of the pathogenesis of egg drop syndrome in laying hens. Avian Pathol. 17:653— 666. 1988.

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20 HEMORRHAGIC ENTERITIS OF TURKEYS MARBLE SPLEEN DISEASE OF PHEASANTS F. William Pierson and Scott D. Fitzgerald SUMMARY. Hemorrhagic enteritis of turkeys and marble spleen disease of pheasants are caused by viruses of the family Adenoviridae, genus Siadenovirus. They are serologically distinct from other avian adenoviruses but are indistinguishable from one another as determined by agar gel immunodiffusion (AGID) test. Slight differences appear at the DNA level and clinical manifestations of disease vary with the strain of virus and the host infected. Most gallinaceous birds are susceptible to infection, but disease is of primary importance in turkeys and pheasants. Agent Identification. Diagnosis can usually be made on the basis of clinical signs, lesions, histopathology, and the demonstration of viral antigen or DNA in splenic tissue by AGID and PCR respectively. Serologic Detection in the Host. Analysis of acute and convalescent sera by AGED or enzyme linked immunosorbent assay may also be of diagnostic value.

INTRODUCTION

Gross Lesions Turkeys that have died from HE are pale due to loss of blood but are otherwise in good flesh and often have feed in the crop. The anterior small intestine is congested and the lumen is often filled with blood. The intestinal mucosa especially at the level of the duodenum has hemorrhage with progression from ecchymoses to petechiae, proximal to distal. In such cases the spleen may appear small and pale but in moribund birds killed prior to extensive blood loss, splenic enlargement with mottling is typically observed (22,36,39). In pheasants, splenic enlargement with mottling, lung congestion and edema are the hallmark lesions. Gastrointestinal lesions are absent (5).

Hemorrhagic enteritis (HE) is an acute viral disease of turkeys, 4 wk of age and older caused by a virus of the family Adenoviridae, genus Siadenovirus, i.e., Turkey Hemorrhagic Enteritis Virus (THEV) (8) In its most severe form the disease is characterized by depression, bloody droppings, and death. Survivors as well as those with milder forms of the disease, often develop secondary coliform infections as a result of transient immunosuppression. This is believed to be the result of viral replication within IgM bearing Blymphocytes and macrophages as well as cytokine mediated apoptosis of bystander cells (40). Gastrointestinal lesion formation is likewise considered to be immune mediated (40). Prevention of the gastrointestinal form of the disease may be accomplished through the use of avirulent live-virus vaccines (14,18). Marble spleen disease (MSD) is a condition affecting captivereared pheasants between 3-8 mo of age and is caused by a virus serologically indistinguishable from THEV (11,28). However, MSD virus (MSDV) primarily causes pulmonary edema and death by asphyxiation in affected pheasants. Mottled enlarged spleens are also characteristic (20). Like HE, fulminate MSD is thought to be an immune mediated disease and may be prevented by vaccination (15). Both THEV and MSD have been reviewed extensively in the literature (20,37).

Histopathology In cases of HE or MSD splenic tissue that has been fixed in 10% neutral buffered formalin and stained with hematoxylin and eosin reveals numerous distinct microscopic lesions. These include lymphoid follicular depletion due to virus-induced apoptosis and necrosis as well as prominent hyperplasia of large lymphoreticular cells within the white pulp, many of which contain eosinophilic intranuclear inclusions (23,29,41,40). Intestinal samples may be of confirmative value if preserved in Zenker's fixative within a few minutes of death and stained with hematoxylin and eosin. Characteristic histopathologic changes include severe congestion of the mucosa, degeneration and detachment of the villus epithelium, and hemorrhage within the lamina propria (41). Occasional intranuclear inclusions may be seen in lymphocytes and macrophages of the lamina propria, in mononuclear cells located within the fiver, bone marrow, peripheral blood, lung, pancreas, brain, and renal tubular epithelium (26,29,41,45). Microscopically the splenic lesions seen with MSD are identical to those of HE. Lung lesions in pheasants that succumb to MSD are consistent with the congestion and edema observed grossly (21). Intranuclear inclusions are occasionally found in mononuclear cells in the lung, liver, proventriculus, bursa of Fabricius, and bone marrow (21).

CLINICAL DISEASE Signs The clinical signs of the gastrointestinal form of HE usually appear 3-4 days after exposure to the virus and include depression, pallor, bloody diarrhea, prostration, and death. Dark red to black gastrointestinal contents may be expressed from the cloaca of dead or moribund birds following application of moderate pressure to the abdomen. Clinically affected birds usually die within 24 hr or recover completely. The disease spreads quickly through susceptible flocks and usually subsides within 6-10 days of onset. The morbidity typically approaches 100% but mortality may range from less than 1 % to as high as 60% depending on the viral pathotype (13,24). Due to the immunosuppressive nature of the virus a second rise in mortality commonly occurs approximately 2 wk after the first due to secondary infections with E. coli, Staphylococcus sp. or other agents (36, 42, 49). The progression of clinical signs associated with MSD in pheasants is also rapid and includes depression, weakness, nasal discharge, dyspnea, and death. Gastrointestinal involvement is not apparent. Mortality usually ranges between 2 and 3% but may reach as high as 20% over the course of approximately 10 days (5,15,33).

SAMPLE COLLECTION THEV can be recovered from the intestinal contents or spleens of dead or moribund turkeys. However for both THEV and MSDV mottled enlarged spleens are by far the best source (12). To obtain useful quantities of virus the capsule of the spleen should be removed and the parenchyma pressed through a syringe to disrupt the reticular architecture. This material should be diluted with an equal volume of 10 mM phosphate buffered saline (pH 7.4), mixed, frozen and thawed 2-3 times, and centrifuged. The resulting supernatant may be used as a test antigen in immunoprecipitation reactions or for further propagation in susceptible birds and cell 90

Chapter 20

culture. The virus is extremely stable and can be maintained for extended periods by freezing or lyophilization. Infectivity is destroyed by drying or application of disinfectants containing phenol or sodium hypochlorite (9,10).

Hemorrhagic Enteritis of Turkeys Marble Spleen Disease of Pheasants

be divided into smaller aliquots, refrigerated, and re-liquefied by heating in a microwave or boiling water prior to the pouring of test plates. Disposable immunodiffusion plates that hold approximately 15 ml of gel or comparable products are commonly used. Wells should be 5 mm in diameter and hold approximately 50 ml. Unknown test material and known positive control antigen should be placed in alternating wells around a central well containing positive control antiserum. Plates should be incubated in a humidified chamber at room temperature and read at 24 and 48 hr. Bands of precipitation showing lines of identity with known positive antigen are indicative of THEV or MSDV infection. Quantification of THEV/MSDV antigen. An antigen capture ELISA for HEV has been described in the literature (47). This technique enables more precise quantitation of viral antigen in tissue extracts than does AGID. Competitive PCR may also be used for quantification in tissue based on the DNA copy number (2). Immunohistochemistry. As an adjunct to routine histopathology, immunofluorescent and immunoperoxidase staining techniques have been described that allow accurate identification of THEV or MSDV intranuclear inclusions in fixed tissues (19,21,41).

PREFERRED CULTURE MEDIA AND SUBSTRATES.

THEV and MSDV can be propagated in susceptible turkeys that are 4 wk-of-age or older. However 5-6 wk of age is generally the best time for infection due to the natural decline of maternal antibody (17). Birds may be inoculated intravenously or orally with about 0.25 ml of a IO’2 dilution of supernatant prepared as previously described. Removal of spleens should be performed 3-4 days after intravenous inoculation or 5-6 days after oral inoculation (12). At these times the spleens will be grossly mottled and enlarged. Successful in vitro propagation of THEV and MSDV can be accomplished using MDTC RP19 cells (American Type Culture Collection, Rockville, MD), a turkey lymphoblastoid cell line derived from a Marek’s disease virus-induced tumor (34) or in leukocytes isolated from the peripheral blood of THEV negative turkeys (46). AGENT IDENTIFICATION

Molecular Identification The THEV/MSDV genome is small in comparison to other members of the family Adenoviridae i.e., 26.3 kb in length (38). This in addition to the presence of unique open reading frames and relatively high A+T content, formally establishes THEV and MSDV as being members of the genus Siadenovirus (7,8,38). Techniques for determining the presence of viral DNA in tissues using both in situ DNA hybridization and polymerase chain reactions (PCR) have been described (2,3,25,43). Due to its enhanced sensitivity PCR may be useful in cases in which the suspicion of HEV infection is high but presence of viral antigen cannot be confirmed by AGID or antigen capture ELISA. A suitable technique involves the amplification of a 273 bp region of the hexon gene. Primers (NHEVF-5’-gtggttcagcagaaagttctt-3’; NHEVR-5’-cagtagactcataagcaactat-3’) are added to the mastermix which contains thermostable Taq DNA polymerase, MgCl2 and dNTPs (Platinum Taq Readymix®, Invitrogen, Grand Island, NY) to a final concentration of 0.2 mM of each primer. Sample DNA extracted from splenic tissue (InstaGene™ Matrix, Bio-Rad Laboratories, Hercules, CA) is then added and the PCR reaction is run (thermocycler program: initial denaturation at 95 C for 2 min; then 35 cycles of 95 C for 15 sec, 58 C for 15 sec, 72 C for 30 sec; and final extension at 72 C for 10 min). PCR products are visualized after agar gel electrophoresis (0.8% TBE-agarose gel, 100V for 1 hour) by staining with ethidium bromide and UV trans­ illumination. Intense bands corresponding with the predicted 273 bp product are considered positive.

Morphology and Physicochemical Properties THEV and MSDV are DNA viruses (27,31). They produce thin bands in sucrose or cesium chloride gradients at a density of 1.321.34 g/ml (4,27). They are non-enveloped icosahedrons having 252 capsomeres of which 240 are non-vertex capsomeres (hexons). The 12 vertex capsomeres (pentons) possess a single fiber (43). Complete virions range in diameter from 70 to 90 nm (27,44).

Biological Properties The viruses causing HE and MSD are believed to be pathotypic variants of the same virus. THEV will cause splenic enlargement and mottling in pheasants without producing significant pulmonary lesions (15). Similarly MSDV infection in turkeys is characterized by splenic enlargement and mottling without gastrointestinal involvement (14). The observed variations in clinical disease are probably as much the result of differences in the host immune response i.e., target shock organ (37,40) as they are differences between viral strains. Other variants exist that produce graded intermediate clinical responses in infected birds (14). Mottling and enlargement of the spleen have also been observed in chickens infected with a virus serologically indistinguishable from THEV and MSDV (16). The disease has been referred to as avian adenovirus group Π splenomegaly (AAS) of chickens but probably only represents the expression of THEV/MSDV in a different gallinaceous host. Except for occasional condemnation due to suspicion of Marek’s disease or lymphoid leukosis, AAS appears to be of little economic significance (16).

SEROLOGICAL DETECTION IN THE HOST

Antigen Detection Agar Gel Immunodiffusion (AGID) Test for Antigen. Isolates of THEV and MSDV are serologically indistinguishable from one another. However an AGID may be used to differentiate HEV and MSDV from other pathogens (12,13). Viral antigen may be obtained from the spleens of suspect cases as described in the section on sample collection. Positive control antigen may be prepared as described in the section on preferred culture media and substrates. Positive control antiserum may be obtained 4 wk post-inoculation. Immunodiffusion gels may be prepared in the following manner. Thirty-two (32.0) grams sodium chloride, 0.8 g sodium azide, and 3.2 g agarose are added to 400 ml distilled water in a 1000-ml Erlenmeyer flask. The mixture is then stirred on a hot plate until boiling, at which time it should clarify. The liquefied gel can then

Agar Gel Immunodiffusion Test for Antibody Anti-HEV or anti-MSDV antibodies can be detected in sera as early as 2-3 wk post-infection by AGID (12,13,30). The assay is identical to that described in the section on agent identification with the exception that unknown test sera should be alternated with known positive control sera and placed in opposition to positive control (splenic) antigen. Testing of both acute and convalescent sera should be performed. Enzyme-Linked Immunosorbent Assay (ELISA) Enzyme-linked immunosorbent assays have been developed for the detection of THEV/MSDV antibody (6,35,47) and are available commercially (Synbiotics Corporation, San Diego, CA). These tests are more sensitive than AGID and may be useful in the quantitation of low levels of antibody such as that found in young 91

F. William Pierson and Scott D. Fitzgerald

16. Domermuth, C. H., J. R. Harris, W. B. Gross, and R. T. DuBose. A naturally occurring infection of chickens with a hemorrhagic enteritis/marble spleen disease type of virus. Avian Dis. 23:479-484.1979. 17. Fadly, A. M, and K. Nazerian. Hemorrhagic enteritis of turkeys: influence of maternal antibody and age at exposure. Avian Dis. 33:778-86. 1989. 18. Fadly, A Μ, K. Nazerian, K. Nagaraja, and G. Below. Field vaccination against hemorrhagic enteritis of turkeys by a cell-culture livevirus vaccine. Avian Dis. 29:768-777. 1985. 19. Fasina, S. O., and J. Fabricant. In vitro studies of hemorrhagic enteritis virus with immunofluorescent antibody technique. Avian Dis. 26:150157.1982. 20. Fitzgerald, S. D., and W. M. Reed. A review of marble spleen disease of ring-necked pheasants. J. Wildl. Dis. 25:455-461. 1989. 21. Fitzgerald, S. D., W. M Reed, and T. Bumstein. Detection of type Π avian adenoviral antigen in tissue sections using immunohistochemical staining. Avian Dis. 36:341-347. 1992. 22. Gross, W. B. Lesions of hemorrhagic enteritis. Avian Dis. 11:684-693. 1967. 23. Gross, W. B., and C. H. Domermuth. Spleen lesions of hemorrhagic enteritis of turkeys. Avian Dis. 20:455-466. 1976. 24. Gross, W. B., and W. E. C. Moore. Hemorrhagic enteritis of turkeys. Avian Dis. 11:296-307. 1967. 25. Hess, M, R. Raue, and Η. M Hafez. PCR for specific detection of haemorrhagic enteritis virus of turkeys, an avian adenovirus. J. Virol. Methods 81:199-203. 1999. 26. Hussain, I., C. U. Choi, B. S. Rings, D. P. Shaw, and K. V. Nagaraja. Pathogenesis of hemorrhagic enteritis virus infection in turkeys. Zentralbl. Veterinarmed. 40:715-726.1993. 27. litis, J. P., and S. B. Daniels. Adenovirus of ringnecked pheasants: purification and partial characterization of marble spleen disease virus. Infect. Immun. 16:701-705. 1977. 28. litis, J. P., R. M Jakowski, and D. S. Wyand. Transmission of marble spleen disease in turkeys and pheasants. Am J. Vet. Res. 36:97-101. 1975. 29. Itakura, C., and H. C. Carlson. Pathology of spontaneous hemorrhagic enteritis of turkeys. Can. J. Comp. Med. 39:310-315.1975. 30. Jakowski, R. M, and D. S. Wyand. Marble spleen disease in ring­ necked pheasants: demonstration of agar gel precipitin antibody in pheasants from an infected flock. J. Wildl. Dis. 8:261-263. 1972. 31. Jucker, M T„ J. R. McQuiston, J. V. van den Hurk, S. M, Boyle, and F. W. Pierson. Characterization of the haemorrhagic enteritis virus genome and the sequence of the putative penton base and core protein genes. J. Gen. Virol. 77:469-479. 1996. 32. MacDougald, L. R. Coccidiosis. In: Diseases of poultry, 11th ed. Y.M Saif., H.J. Barnes, J.R. Glisson, A.M Fadly. L.R. McDougald, and D.E. Swayne, eds. Iowa State University Press, Ames, Iowa. pp. 988. 2003. 33. Mayeda, B., G. B. West, A A. Bickford, and B. R. Cho. Marble spleen disease in pen-raised pheasants in California. Proc. Amer. Assoc. Vet. Lab. Diagn. 25:261-270. 1982. 34. Nazerian, K., and A. Fadly. Propagation of virulent and avirulent turkey hemorrhagic enteritis virus in cell culture. Avian Dis. 26:816827.1982. 35. Nazerian, K., and A M Fadly. Further studies on in vitro and in vivo assays of hemorrhagic enteritis virus (HEV). Avian Dis. 31:234-240. 1987. 36. Pierson, F. W., V. D. Barta, D. Boyd, and W. S. Thompson. Exposure to multiple infectious agents and the development of colibacillosis in turkeys. J. Appl. Poult. Res. 5:347-357.1996. 37. Pierson, F. W., and S. D. Fitzgerald. Hemorrhagic enteritis and related infections. In: Diseases of poultry, 11th ed. Y.M Saif, H.J. Barnes, J.R. Glisson, A.M. Fadly, L.R. McDougald, and D.E. Swayne, eds. Iowa State University Press, Ames, Iowa. pp. 237-247. 2003. 38. Pitcovski, J., M Mualem, Z. Rei-Koren, S. Krispel, E. Shmueli, Y. Peretz, B. Gutter, G.E. Gallili, A. Michael, and D. Goldberg. The complete DNA sequence on genome organization of the avian adenovirus, hemorrhagic enteritis virus. Virol 249:307-315. 1998. 39 Pomeroy, B. S., and R. Fenstermacher. Hemorrhagic enteritis in turkeys. Poult. Sci. 16:378-382. 1937. 40. Rautenschlien, S., and J. M Sharma. Immunopathogenesis of haemorrhagic enteritis virus (HEV) in turkeys. Devel. Comp. Immuno. 24:237-246. 2000. 41. Saunders, G. K., F. W. Pierson, and J. V. van den Hurk. Haemorrhagic enteritis virus infection in turkeys: a comparison of virulent and avirulent virus infections and a proposed pathogenesis. Avian Pathol. 22:47-58. 1993.

birds prior to and shortly after vaccination. In older birds with high levels of antibody e.g., breeders or market age turkeys, additional dilution of sera may be required to bring optical densities within readable limits. In most cases multiplication of the sample to positive ratio by the appropriate dilution factor prior to the calculation of the log titer will enable comparison with younger birds. DIFFERENTIATION FROM CLOSELY RELATED AGENTS

In turkeys the presence of mottled enlarged spleens in the absence of detectable HEV antigen or intestinal bleeding should arouse suspicion of other diseases such as reticuloendotheliosis (see Chapter 35) or lymphoproliferative disease (1). Splenic enlargement may also be indicative of a generalized septicemia or other acute conditions (37, see Chapters 3, 4, and 9). Congestion and hemorrhage in the proximal small intestine should also prompt consideration of coccidiosis (Eimeria meleagrimidis) (32). In pheasants mortality associated with asphyxia, dyspnea, pulmonary edema, and mottled enlarged spleens may be considered pathognomonic for MSD. ACKNOWLEGEMENT

The authors are greatly indebted to Drs. C. H. Domermuth and W. B. Gross for their contributions to earlier editions of this chapter. REFERENCES

1. Biggs, P. M Lymphoproliferative disease of turkeys. In: Diseases of poultry, 10th ed B. W Calnek. H. J. Barnes, C. W. Beard, L R. McDougald, Y. M Saif, eds. Iowa State University Press, Ames, Iowa. pp. 485-488. 1997. 2. Beach, N., C. Cardona, R.B. Duncan, Jr., D. Wise, and F. W. Pierson. Competitive PCR for the quantification of hemorrhagic enteritis virus. In: Proceedings of the 73rd Northeastern Conference on Avian Diseases, College Park, Maryland 2001. 3. Beach, N., C. Cardona, D. Wise, R. Duncan, and F. W. Pierson. Latency of hemorrhagic enteritis virus as determined by nested PCR. In: Proceedings of the 74th Northeastern Conference on Avian Diseases, Mystic, Connecticut. 2002. 4. Carlson, H. C., F. AI-Sheikhly, J R. Pettit, and G. L. Seawright. Virus particles in spleens and intestines of turkeys with hemorrhagic enteritis. Avian Dis. 18:67-73. 1974. 5. Carlson, H. C., J. R. Pettit, R. V. Hemsley, and W. R. Mitchell. Marble spleen disease of pheasants in Ontario. Can. J. Comp. Med. 37:281-286. 1973. 6. Davidson, I., A. Aronovici, Y. Weisman, and M Malkinson. Enzyme immunoassay studies on the serological response of turkeys to hemorrhagic enteritis virus. Avian Dis. 29:43-52. 1985. 7. Davison, A J., and B. Harrach. Siadenovirus. In: The springer index of viruses. C.A. Tidona and G. Darai, eds.: Springer-Verlag, New York, New York, pp.29-33. 2002. 8. Davison, A.J., M Benko, and B. Harrach. Genetic Content and Evolution of Adenoviruses. J. Gen. Virol. 84:2895-2908. 2003. 9. Domermuth, C. H., and W. B. Gross. Effect of disinfectants and drying on the virus of hemorrhagic enteritis of turkeys. Avian Dis. 15:94-97. 1971. 10. Domermuth, C. H., and W. B. Gross. Effect of chlorine on the virus of hemorrhagic enteritis of turkeys. Avian Dis. 16:952-953. 1972. 11. Domermuth, C. H., and W. B. Gross. Hemorrhagic enteritis of turkeys: antiserum efficacy, preparation and use. Avian Dis. 19:657-665. 1975. 12. Domermuth, C. H., W. B. Gross, R. T. DuBose, C. S. Douglass, and C. B. Reubush, Jr. Agar gel diffusion precipitin test for hemorrhagic enteritis of turkeys. Avian Dis. 16:852-857. 1972. 13. Domermuth, C. H., W. B. Gross, and R. T. DuBose. Microimmunodiffusion test for hemorrhagic enteritis of turkeys. Avian Dis. 17:439-444. 1973. 14. Domermuth, C. H., W. B. Gross, C. S. Douglass, R. T. DuBose, J. R. Harris, and R. B. Davis. Vaccination for hemorrhagic enteritis of turkeys. Avian Dis. 21:557-565. 1977. 15. Domermuth, C. H., W. B. Gross, L. D. Schwartz, E. T. Mallinson, and R. Britt. Vaccination of ring-necked pheasant for marble spleen disease. Avian Dis. 23:30-38. 1979.

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46. van den Hurk, J. V. Propagation of hemorrhagic enteritis virus in normal (non-tumor derived) cell culture. J. Am. Vet. Med. Assoc. 187:307. 1985. 47. van den Hurk, J. V. Quantitation of hemorrhagic enteritis virus antigen and antibody using enzyme-linked immunosorbent assays. Avian Dis. 30:662-671. 1986. 48. van den Hurk, J. V. Characterization of the structural proteins of hemorrhagic enteritis virus. Arch. Virol. 126:195-213. 1992. 49. van den Hurk, J. V., B. J. Allan, C. Riddell, T. Watts, and A. A Potter. Effect of infection with hemorrhagic enteritis virus on susceptibility of turkeys to Escherichia coli. Avian Dis. 38:708-716. 1994.

42. Sponenberg, D. P., C. H. Domermuth, and C. T. Larsen. Field outbreaks of colibacillosis of turkeys associated with hemorrhagic enteritis virus. Avian Dis. 29:838-842. 1985. 43. Suresh, M., and J. M Sharma. Pathogenesis of type II avian adenovirus infection in turkeys: in vivo immune cell tropism and tissue distribution of the virus. J. Virol. 70:30-36. 1996. 44. Tolin, S. A., and C. H. Domermuth. Hemorrhagic enteritis of turkeys. Electron microscopy of the causal virus. Avian Dis. 19:118-125. 1975. 45. Trampel, D. W., C. U. Meteyer, and A. A Bickford. Hemorrhagic enteritis virus inclusions in turkey renal tubular epithelium. Avian Dis. 36:1086-1091. 1992.

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21 INFECTIOUS LARYNGOTRACHEITIS Deoki N. Tripathy and Maricarmen Garcia

SUMMARY. Infectious laryngotracheitis (ILT) is either an acute, highly contagious herpesvirus infection of chickens, characterized by severe dyspnea, coughing, and rales; or a subacute disease with lacrimation, tracheitis, conjunctivitis, and mild rales. ILT has been reported from most of the intensive poultry-rearing areas of the United States and many other countries. Mortality varies and may reach 50% in adult birds, usually due to occlusion of the trachea by hemorrhage or exudate. Following recovery, some birds remain carriers for variable periods and become a source of infection for susceptible birds. Diagnosis during the acute stage of infection is usually made by clinical signs and demonstration of intranuclear inclusion bodies in the infected tracheal epithelium. Agent Identification. Infectious laryngotracheitis is routinely diagnosed by histopathologic examination of tracheal lesions characterized by the presence of intranuclear inclusion bodies. In addition, viral particles exhibiting typical herpesvirus morphology can be detected by negative staining of lesion suspensions or by direct examination of ultrathin sections of affected tissues using transmission electron microscopy. The virus is isolated by inoculation of chorioallantoic membrane (CAM) of 9 to 12-day-old developing chicken embryos. Lesions develop on the CAM 5 to 7 days post-infection. Etiology is confirmed by histopathologic examination of the CAM for intranuclear inclusion bodies or electron microscopy for herpesvirus morphology. Isolated viruses can be evaluated for pathogenicity in susceptible hosts, for example, chickens (development of clinical signs and tracheal lesions), or for growth and pathogenicity of avian cell culture (cytopathic effect and plaque formation). Genomic characterization of virus isolates can be based on restriction fragment length polymorphism (RFLP), in which the DNA profiles are compared after restriction enzyme digestion and subsequent agarose gel electrophoresis. Although cloned genomic fragments have been used as diagnostic probes, the use of polymerase chain reaction (PCR) for the amplification of a specific portion of the genome have been used more frequently as a rapid diagnostic method for detection of viral nucleic acid in experimentally and naturally infected birds. Although PCR has not been used as a routine diagnostic tool, PCR-RFLP procedures has been widely utilized as an epidemiological tool to differentiate among viral isolates. Other rapid diagnostic methods utilized are the immunoperoxidase staining, indirect fluorescent antibody, and the agar-gel immunodiffusion (AGID) test, all utilized to detect the viral antigen in affected tissues. Serologic Detection in the Host. Serologic detection of infection may be important in experimental studies and in measuring immune response following vaccination. Antibody response can be measured by AGID tests, the immunoperoxidase (IP) test, and the enzyme-linked immunosorbent assays (ELISA). The immunoperoxidase/indirect fluorescent antibody and AGID tests can also be used to detect the antigen in affected tissues. Antigenic differences among isolates can be determined by host pathogenicity and virus neutralization. Vaccination with modified live strains of low virulence is practiced in endemic areas and on farms where a specific diagnosis has been made. Vaccination in the face of an outbreak shortens the course of the disease.

may be detectable in some cases. In the milder forms of the disease, only a few birds in a flock show classical respiratory signs, and mortality is low. In such cases, the clinical signs are characterized by mild tracheatis, swollen sinus and mild conjunctivitis (32, 37). Turkeys and pheasants have a low degree of susceptibility to ILTV (18).

INTRODUCTION

Infectious laryngotracheitis (ILT) is an acute respiratory disease of chickens caused by an alphaherpesvirus. ILT has been recognized in many countries and is an economically important disease of chickens (18). In areas with large concentrations of poultry, ILT is responsible for losses due to lowered egg production and mortality. As a member of the alpha-herpesvirus family the ILT virus (ILTV) has the ability to establish latency and to persist in recovered chickens for variable periods after recovery from the disease. Thus, the carrier state is of considerable epizootiological significance, as carrier birds may shed the reactivated virus and may be responsible for new outbreaks of the disease.

SAMPLE COLLECTION

Tracheal exudates, tracheal scrapings and lung should be collected for virus isolation in the early acute phase of the disease. A 10% suspension is made of the tissue in sterile normal saline, Hanks balanced salt solution, or broth by grinding the tissue either with sterile fine sand or 60-mesh aluminum oxide with a sterile mortar and pestle or in a glass grinder. Alternatively tracheal scrapings can be collected with a sterile scalpel and resuspended in any suitable sterile transport media. The suspension is centrifuged at 700 x g for 10 min, and antibiotics (1000 IU/ml penicillin and 1 g/ml of dihydrostreptomycin) are added. The supernatant fluid is held at room temperature for 30-50 min before inoculation into the culture substrate. Tracheal tissue from early acute cases may be collected in Bouin’s or Zenker’s fixatives for demonstration of intranuclear inclusion bodies by histopathologic examination. Serum samples may be collected to measure antibody responses by virus-neutralization (VN), agar-gel immunodiffusion (AGID), enzyme-linked immunosorbent assay (ELISA), or fluorescent antibody (FA) tests to detect specific antibodies.

CLINICAL DISEASE Infection with ILTV is characterized by anorexia, depression, and severe respiratory distress, with coughing, sneezing, gurgling, gasping, and rales. The neck is often extended during inspiratory efforts, because the trachea is often partially occluded, and bloody, mucous exudate may be expelled during violent expectorating efforts. Conjunctivitis has been characteristic of more recent outbreaks Although ILTV causes a highly localized infection involving mainly the trachea and the conjunctiva, extratracheal spread of the virus to the lung, trigeminal ganglion and brain has also been reported. In severe forms of the disease, mortality is as high as 50% or more, with significant lesions consisting of hemorrhages in the trachea and larynx. The tracheal lumen is often filled with blood clots, mucus, caseous yellowish exudate, or a tracheal plug, which may cause death from asphyxia. Intranuclear inclusion bodies are detectable in the early stage of the disease on histopathologic examination of the infected trachea. Air sacculitis and pneumonitis 94

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Camoys solution and stained either by the Feulgen method or with acridine orange) inclusion bodies in a few multinucleate giant cell nuclei are observed 12 hr after infection. Degenerate and detached cells are seen from 36 to 72 hr, and degeneration may be complete by 72 hr PI. Adult chicken kidney culture, CEK cells, and the LMH cell line have been used as a plaque system for quantitative assay of ILTV. Plaques of 1 to 2 mm in diameter develop by 4 to 5 days PI (35, 39).

PREFERRED CULTURE MEDIA AND SUBSTRATES

Routinely ILTV can be isolated from tracheal exudates, tracheal scrapings or lung tissue suspensions by inoculation of developing embryonated eggs, the trachea and infraorbital sinuses of susceptible chickens, or inoculation of chick embryo liver (CEL) and chicken embryo kidney (CEK) monolayer cultures and virus isolation from tracheas of latently infected birds has been possible using the tracheal organ culture system (18).

AGENT IDENTIFICATION Embryo Inoculation Chicken embryos, 9 to 12 day-old, from specific-pathogen-free flocks are inoculated on the chorioallantoic membrane (CAM) with a 0.1 to 0.2-ml suspension of tracheal exudate or tracheal and lung tissues. Inoculated embiyos are incubated at 37 C. As early as 3 days post-inoculation (PI), plaques can be observed on the CAM that have opaque edges and depressed central areas of necrosis. The plaques result from proliferation and necrosis of the affected cells and may vary from a few scattered foci to large numbers. Some embryos die between 2 to 8 days PI, and the size of surviving embryos is often reduced.

Infectious laryngotracheitis (ILT) is routinely diagnosed by histopathologic examination of tracheal lesions for the presence of intranuclear inclusion bodies. In addition, viral particles exhibiting typical herpesvirus morphology can be detected by negative staining of suspensions from affected tissues or by direct examination of ultrathin sections with electron microscopy. On inoculated CAM the etiology is confirmed by histopathologic or ultrastructural examination of the CAM lesions for intranuclear inclusion bodies or viral particles with herpesvirus morphology. Isolated viruses can be evaluated for pathogenicity in susceptible hosts, for example, in chickens (development of clinical signs and tracheal lesions), or for susceptibility of avian cell culture (cytopathic effect and plaque formation). A slide-smear technique for diagnosis of ILT has been described (7). Smears from scrapings of laryngeal mucosa (or conjunctiva in the case of conjunctivitis) are fixed for 3-5 min in absolute methyl alcohol and stained with Giemsa (1 drop of stock per ml of distilled water) for 2 hr at 37 C or overnight at room temperature. Stained smears are washed in tap water and differentiated in absolute methyl alcohol, using three to five quick dipping motions, until the smear has a magenta cast. The smear is washed in tap water, air­ dried, and examined. Intranuclear inclusion bodies of ILTV are detectable in the early acute stages of disease. Tracheal impression smears stained by Giemsa in early stages of the disease also reveal intranuclear inclusion bodies. The inclusions appear purple with a pinkish cast and are surrounded by a halo. Inclusions can be observed in conjunctival smears in cases with conjunctivitis. Sevoian (38) described a quick method for diagnosis of ILT. This method uses simultaneous fixation and dehydration and demonstrates intranuclear inclusion bodies in less than 3 hr. In another histologic diagnostic method tracheas and affected tissues are embedded in wax medium for sectioning and satisfactory slides are prepared in 3 hr (33). Direct electron microscopy can be used to identify virions of typical herpesvirus morphology in lysed cells from tracheal scrapings of infected birds. Various serologic tests (e.g., FA, immunoperoxidase [IP], and AGID), described later in the chapter, can be used to detect the ILTV antigen in the specimen.

Chicken Inoculation Susceptible chickens can be inoculated by tracheal, infraorbital sinus, or aerosol methods with a suspension of the specimen. Under experimental conditions the severity of respiratory signs and mortality depends on virulence of the viral strain, susceptibility and age of the birds, and the route of inoculation. The virus multiplies primarily in the trachea and can be isolated 2 to 7 days after intratracheal inoculation. Virus titer is at a maximum 3 to 4 days PI. Chickens infected with virulent strains usually develop signs of respiratory tract infection by day 3 PI. Most chickens show typical signs of the disease by days 4 and 5. Mild respiratory disease with mild, focal, non-suppurative pneumonitis and air sacculitis may be observed in some birds. Gross changes in birds infected experimentally with virulent ILTV by the aerosol method consist of severe hemorrhagic laryngitis and tracheitis by day 3 PI, with extension to the syrinx and bronchi by day 4. A yellowish, diphtheritic membrane lining the trachea and larynx is observed at about day 6 PI and may be found as a plug in the laiynx and upper trachea by day 8. The tracheal exudate is expelled by day 10. Gross lesions of the lungs and air sacs are observed in some experimental chickens between days 3 and 7 PI and occasionally for longer periods. During the incubation stage in experimentally infected, susceptible birds, isolated areas of epithelial hyperplasia appearing as giant cell syncytia occur in the trachea and laiynx by 48 hr PI. Intranuclear inclusion bodies are detectable in the syncytia. Necrotizing tracheitis with sloughing of the epithelium in cell groups with intranuclear inclusion bodies is seen by day 3 PI. Inclusions are usually observed from 4 to 6 days PI. At 5 to 7 days PI, the lesion progresses to extensive necrosis of the epithelium, and inclusions are visible less often. In recovered birds, the epithelium regenerates to normal appearance by day 12 (34). Pneumonitis and airsacculitis are often present during the acute stage, and giant cell syncytia containing intranuclear inclusion bodies are observed in the air sac walls and lungs.

Fluorescent Antibody (FA) The viral antigen can be detected by direct and indirect FA in tracheal smears or frozen tracheal sections during the early acute phase of the disease (2 to 8 days after exposure) 22). The FA test is a useful rapid assay. Direct FA on smears of tracheal scrapings has been evaluated on experimentally and naturally infected birds. On naturally infected birds, viral antigens were detected from 2 to 14 days PI, while characteristic ILTV lesions were detected by histopathological examination were detected from day 3 to 10 PI (43). In naturally infected birds, FA and histopathological examination were equally satisfactory in the detection of infected birds (15).

Cell Culture Chick embryo liver (CEL) and chicken embryo kidney (CEK) cells are suitable for isolation of ILTV (23). A chicken liver tumor cell line, LMH, has also been used for propagation of ILTV (39). Early cytopathic changes in the CEK cells after inoculation with ILTV consist of the development of numerous multinucleate giant cells that grow, coalesce, and finally undergo degeneration with continued incubation. Large, basophilic (methanol-fixed and stained by the May-Griinwald-Giemsa method), DNA-positive (fixed in

Immunoperoxidase (IP) An indirect IP method with monoclonal antibody has been used for detection of ILTV antigen in frozen tracheal sections (17). In a recent experimental study (2), IP was more sensitive than other 95

Deoki N. Tripathy and Maricarmen Garcia

diagnostic tests when histopathology, FA, hybridization were compared.

IP, PCR,

and

sequences (10, 14). In these reports outbreak related isolates collected before the use of vaccination were easily differentiated from vaccines strains. However, outbreak related isolates obtained after the implementation of vaccination were identical to the currently utilized vaccines. A PCR-RFLP analysis of the viral glycoprotein E (gE) gene demonstrated that different viral subpopulations are present in the chicken embryo origin (CEO) ILTV vaccine preparations. This assay permitted the identification of vaccine related isolates in as the cause of outbreaks in the US (13). Korean field isolates were differentiated from vaccine strains by PCR-RFLP analysis of the glycoprotein G and thymidine kinase (tk) genes (20). Although vaccine strains and field isolates from different countries were differentiated successfully using different PCR-RFLP none of the reported ILTV DNA differences between vaccine and field isolates has been related to differences in strains pathogenicity at this moment.

Molecular Identification Procedures for detection of ILTV DNA using dot-blot hybridization assays with cloned ILTV DNA fragments labeled with p32 or digoxigenin have been described (12, 26, 29). These procedures demonstrated to be highly sensitive for detection of ILTV in acutely affected and convalescent chickens when detection was no longer possible by virus isolation. With the advent of polymerase chain reaction (PCR) several viral nucleic acid amplification procedures have been described (1, 4, 24, 40, 41, 44, 45). Some PCR have included the use of non-isotopically labeled probes on membrane hybridization (1, 4), or an ELISA format, to enzymatically detect the amplification products (40). Otherwise amplification products are visualized by gel electrophoresis. Nested PCR amplifications, with two sets of primers, have been utilized to increase the sensitivity of detection of ILTV DNA (24). Different samples have been evaluated for the detection of ILTV DNA by PCR. For example, viral DNA has been successfully amplified from ILTV infected cell culture supernatants (40), tracheal scrapings (41, 45), tracheal swabs (44), conjunctival swabs (4), turbinates, trigeminal ganglia (44), and from formalin-fixed, paraffin embedded tracheal tissues (24). The ability of PCR to detect ILTV infected birds has been compared to other diagnostic assays in samples from experimentally (2, 4) and naturally infected birds (45). In tracheas from experimentally infected birds PCR was more effective than virus isolation in the detection of ILTV, particularly during late stages of infection when birds have recovered from clinical signs. In naturally infected birds PCR detected viral DNA from tracheal samples that contained bacteria and other viruses that prevented the isolation of ILTV in cell culture (45). Molecular methods for differentiating ILTV strains have been described. These include restriction fragment length polymorphism (RFLP) analysis of the viral genome (6, 16, 19, 27, 28), reciprocal DNA : DNA hybridization using cloned DNA fragments (31, 42), and RFLP of PCR amplified products (9, 10, 13, 14, 20). RFLP analysis of the viral genome requires large amounts of purified viral DNA, reciprocal hybridization protocols call for in-vitro radiolabelling of viral DNA. Although these procedures are useful in epidemiological studies they are cumbersome not practical for larger epidemiological studies, and not feasible for the daily diagnostic routine. The use of PCR based methods has greatly facilitated the molecular differentiation of ILTV because pure viral DNA is not required, and a large number of isolates from a single outbreak, or from different geographical regions can be analyzed in a timely fashion.

SEROLOGIC DETECTION IN THE HOST Agar-Gel Immunodiffusion Infectious laryngotracheitis virus antigen is prepared from infected CAMs of embryonated eggs by homogenizing membranes that have confluent lesions. Large tissue particles are removed by centrifugation at 1600 x g, and the supernatant is used as an antigen. Tracheal antigen is prepared from a bird that either died of ILT or was euthanized within 1 wk of the onset of clinical signs. The antigen consists of exudate collected from the lumen and walls of the larynx, trachea, and extrapulmonary bronchi. If the exudate is too dry, it is diluted with normal saline. Antigen prepared from either the ILTV-infected CAM or the tracheal exudate is reacted with specific antibody in plates prepared with 1.5% Noble agar and 8% sodium chloride in veronal buffer (pH 7.2). Sodium azide in 1% concentration of thimerosal in 0.01% concentration is added as a preservative. A line or lines of precipitation will develop between the antigen and antibody wells after 24 to 48 hr of incubation at room temperature if the concentration of antigen is adequate in the tracheal exudate. Because the amount of precipitating antigen in the tracheal exudate and the amount of live virus will vary with the period of infection and from bird to bird, AGID should be performed in combination with egg or cell culture inoculation (and tests such as IP, FA, or PCR). A small amount of five virus present in tracheal exudate will infect the CAM of embryonated eggs or cell culture, whereas an inadequate concentration of precipitating antigen will not form a line of precipitation with the antibody. AGID also can be used to detect the antibody responses of recovered birds by reacting the sera with known antigen prepared from ILTV-infected cell cultures or CAM (25). A positive response may occur in 25%-50% of the birds, and occasionally in over 80%. The test is less sensitive than IP, ELISA, FA, and VN. However, it still may be a useful test for detecting antibodies on a flock basis (3).

Strain Variability Naturally occurring strains of ILTV of variable pathogenicity have been isolated from field outbreaks. However, not enough information is available on their characterization. The DNA of virulent and avirulent strains shows high homology by reciprocal DNA-DNA hybridization (30, 42). However, some minor differences can be detected by restriction endonuclease analysis (27) and by PCR RFLP analysis (9, 11, 14, 42). A PCR-RFLP assay developed by Clavijo and Nagy (9) allowed the differentiation of a virulent LT isolate from a low less virulent Canadian isolate. Australian vaccine and a pathogenic field strain were differentiated by the identification of a 467 base pair insertion/deletion in the ICP4 gene non-coding region (42). The 467 bp pair deletion present in the Australian pathogenic field isolate was not detected in other pathogenic field isolates from Taiwan (11). However, Taiwanese isolates as well as isolates from England, Scotland, and the Republic of Ireland were differentiated from vaccine strains by PCR-RFLP analysis of the coding and non-coding ICP4 gene

Virus Neutralization Infectious laryngotracheitis virus or ILTV antibody can be identified by the VN test. Undiluted serum mixed with 10-fold virus dilutions or serially diluted serum mixed with a single concentration of virus is held at room temperature for 1^4 hr before inoculation of 9-to-12-day-old embryonated chicken eggs or CEK cell cultures. Neutralizing antibody is detectable by 1 wk PI (21). A microassay and neutralization test with CEK (35) and CEL cells (5) in microcell culture trays can be used satisfactorily.

Enzyme-Linked Immunosorbent Assay An ELISA tests has been developed to detect serum antibodies against ILTV and is available commercially in United States and Europe. Current ELISA tests utilize antigen prepared from either infected CAM or infected cell cultures. Using antigen prepared 96

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Infectious Laryngotracheitis

16. Guy, J. S., H. J. Bames, L. L. Munger, and L. Rose. Restriction endonuclease analysis of infectious laryngotracheitis viruses: Comparison of modified-live vaccine viruses and North Carolina field isolates. Avian Dis. 33:316-323.1989. 17. Guy, J. S., H. J. Bames, and L. G. Smith. Rapid diagnosis of infectious laryngotracheitis using a monoclonal antibody-based immunoperoxidase procedure. Avian Path. 21:77-86. 1992. 18. Guy, J. S. and T. J. Bagust. Laryngotracheitis. In: Diseases of Poultry, 11th ed. Y. M Saif, ed. Iowa State University Press, Ames, Iowa. pp. 121134.2003. 19. Han, M G. and S. J. Kim. Comparison of virulence and restriction endonuclease cleavage patterns of infectious laryngotracheitis virus isolated in Korea. Avian Pathol. 30: 337 -344. 2001a. 20. Han, M G., and S. J. Kim. Analysis of Korean strains of infectious larymgotracheitis virus by nucleotide sequences and restriction fragment length polymorphism. Veterinary Microbiol. 83:321 - 331. 2001b. 21. Hitchner, S. B., C. A. Shea, and P. G. White. Studies on a serum neutralization test for the diagnosis of laryngotracheitis in chickens. Avian Dis. 2:258-269. 1958. 22. Hitchner, S. B., J. Fabricant, and T. J. Bagust. A fluorescent-antibody study of the pathogenesis of infectious laryngotracheitis. Avian Dis. 21:185— 194. 1977. 23. Hughes, C. S., and R. C. Jones. Comparison of cultural methods for primary isolation of infectious laryngotracheitis virus from field material. Avian Pathol. 17:295-303. 1988. 24. Humberd, J., Riblet, S., Resurreccion, R.S., Brown, T.P., and Garcia, M. Detection of infectious laryngotracheitis in formalin-fixed, paraffin embedded tissues by nested-PCR Avian Dis. 46: 64-74. 2002. 25. Jordan, F. T. W., and R. C. Chubb. The agar gel diffusion technique in the diagnosis of infectious laryngotracheitis (ILT) and its differentiation from fowlpox. Res. Vet. Sci. 3:245-255. 1962. 26. Keam L, York JJ, Sheppard M, Fahey KJ.Detection of infectious laryngotracheitis virus in chickens using a non-radioactive DNA probe. Avian Dis.35:257-262. 1991. 27. Keeler, C. L., Jr., J. W. Hazel, J. E. Hastings, and J. K. Rosenberger. Restriction endonuclease analysis of Delmarva field isolates of infectious laryngotracheitis virus. Avian Dis. 37:418-426. 1993. 28. Keller, L. H., C. E. Benson, S. Davison, andR. J. Eckroade. Differences among restriction endonuclease DNA fingerprints of Pennsylvania field isolates, vaccine strains, and challenge strains of Infectious laryngotracheitis virus. Avian Dis. 36:575-581. 1992. 29. Key, D. W., C. B. Gough, J. B. Derbyshire, and E. Nagy. Development and evaluation of a non-isotopically labeled DNA probe for the diagnosis of infectious laryngotracheitis. Avian Dis. 38:467-474. 1994. 30. Kotiw M, Sheppard M, May JT, Wilks CR. Differentiation between virulent and avirulent strains of infectious laryngotracheitis virus by DNA: DNA hybridization using a cloned DNA marker. Vet Microbiol. 11(4):319-30. 1986 31. Leong, V.Y., J. R. Glisson, R. S. Resurreccion, and I-H. N. Cheng. Infectious laryngotracheitis virus in commercial hens: A serological study based on enzyme - linked immunosorbent assay. Avian Dis.38:304 - 307. 1994. 32. Linares, J. A., A. A. Bickford, G. L. Cooper, B. R. Charlton, and P. R. Woolcock. An of infectious laryngotracheitis virus in California broilers. Avian Dis. 38:188-192. 1994. 33. Pirozok, R. P., C. F. Helmboldt, and E. L. Juhgherr. A rapid histological technique for the diagnosis of infectious avian laryngotracheitis. J. Am. Vet. Med. Assoc. 130:406-407. 1957. 34. Purcell, D. A. Histopathology of infectious laryngotracheitis in fowl infected by an aerosol. J. Comp. Pathol. 81:421^131. 1971. 35. Robertson, G. M, and J. R. Egerton. Micro-assay systems for infectious laryngotracheitis virus. Avian Dis. 21:133-135. 1977. 36. Sander, J. E. and S. G. Thayer. Evaluation of ELISA titers to infectious laryngotracheitis. Avian Dis. 41:429-432.1997. 37. Sellers, H. S., M Garcia, J. R. Glisson, T. P. Brown, J. S. Sander, and J. S. Guy. Mild infectious laryngotracheitis in broilers in the Southeast. Avian Dis. 48:430-436.2004. 38. Sevoian, M A. A quick method for the diagnosis of avian pox and infectious laryngotracheitis. Avian Dis. 4:474-476. 1960. 39. Schnitzlein, W. M, J. Radzevicius, and D. N. Tripathy. Propagation of infectious laryngotracheitis virus in avian liver cell line. Avian Dis. 38:211— 217. 1994. 40. Shirley MW, Kemp DJ, Sheppard M, Fahey KJ. Detection of DNA from infectious laryngotracheitis virus by colourimetric analyses of polymerase chain reactions. J Virol Methods. 30(3):251 -9. 1990.

from either infected CAM or infected cell cultures can detect antibodies in the sera of infected birds as early as 7 days PI (3). The ELISA was shown to be more sensitive than VN in the detection of ILTV serum antibodies (8). ELISA is an ideal test for survey purposes particularly when it has been utilized to evaluate the antibody response of vaccinated flocks (31, 36). Antigen capture ELISA using ILTV polyclonal or monoclonal antibodies and then detecting the capture antigen by enzyme-labeled antibodies has also been utilized as a rapid diagnostic test to detect infected flocks (46).

DIFFERENTIATION FROM CLOSELY RELATED AGENTS Although acute signs of ILT, characterized by coughing, expulsion of blood, and high mortality, can be suggestive of ILT, many signs are similar to those of other respiratory diseases. The diphtheritic form of fowlpox with lesions in the trachea may simulate signs of ILT. Respiratory signs caused by mildly virulent strains of ILTV may be indistinguishable from other respiratory infections, such as mycoplasmosis and infectious bronchitis. Therefore tests such as PCR, FA, IP become relevant confirmatory tests in the diagnosis of ILTV. REFERENCES

1. Abbas, F., J. R. Andreasen, Jr., and M W. Jackwood. Development of a polymerase chain reaction and a nonradioactive DNA probe for infectious laryngotracheitis virus. Avian Dis. 40:56-62. 1996. 2. Abbas, F., and J. R. Andreasen, Jr. Comparison of diagnostic tests for infectious laryngotracheitis. Avian Dis. 40:290-295. 1996. 3. Adair, B. M, D. Todd, E. R. McKillop, and K. Bums. Comparison of serological tests for detection of antibodies to infectious laryngotracheitis virus. Avian Pathol. 14:461-469. 1985. 4. Alexander, H. S., and E. Nagy. Polymerase chain reaction to detect infectious laryngotracheitis virus in conjunctival swabs from experimentally infected chickens. Avian Dis. 41:646-653. 1997. 5. Andreasen J. R, J. Brown, J. R. Glisson and P. Villegas. Reprodicibility of a virus-neutralization test for infectious laryngotracheitis virus. Avian Dis. 34:185-192. 1989. 6. Andreasen, J. R., J. Glisson, Jr., and P. Villegas. Differentiation of vaccine strains and Georgia field isolates of Laryngotracheitis virus by theis restriction endonuclease. Avian Dis. 34:646-656. 1990. 7. Armstrong, W. H. A slide smear technique for the diagnosis of laryngotracheitis. Avian Dis. 3:80-84. 1959. 8. Bauer, B.,J. E.Lohr and E. F. Kaleta. Comparison of commercial ELISA test kits from Australia and the USA with the serum neutralization test in cell cultures for the detection of antibodies to the infectious laryngotracheitis virus of chickens. Avian Pathology. 28:65-72. 1999. 9. Clavijo, A., and E. Nagy. Differentiation of laryngotracheitis virus strains by polymerase chain reaction. Avian Dis. 41:241-246. 1997. 10. Chang, P. C., Y. L. Lee, J. H. Shien, and Η. K. Shien. Rapid differentiation of vaccine strains and field isolates of infectious laryngotracheitis virus by restriction fragment length polymorphism of PCR products. J. Virol. Methods. 66:179-186. 1997. 11. Chang, P. C.,H. K.Shieh, J. H. Shien, and S. W. Kang. A homopolymer stretch composed of variable numbers of cytidine residues in the terminal repeats of infectious laryngotracheitis virus. Avian Dis. 44:125-131.2000. 12. Fatunmbi, Ο. O., W. M Reed, D. L. Schwartz, and D. N. Tripathy. Dual infection of chickens with pox and infectious laryngotracheitis (ILT) confirmed with specific pox and ILT DNA dot-blot hybridization assays. Avian Dis. 39:925-930. 1995. 13. Garcia, M, and S. M Riblet. Characterization of Infectious laryngotracheitis virus (ILTV) vaccine strains and field isolates: demonstration of viral sub-populations within vaccine preparations. Avian Dis. 45:558-566. 2001. 14. Graham, D. A., I. E. Mclaren, V. Calvert, D. Torrens, and B. M Meeham. RFLP analysis of recent Northern Ireland isolates of infectious laryngotracheitis: comparison with vaccine virus and field isolates from England, Scotland and Republic of Ireland. Avian Pathol. 29:57-62. 2000. 15. Goodwin, M. A, M A. Smeltzer, J. Brown, R. S. Resurreccion, and T. G. Dickson. Comparison of histopathology to the direct immunofluorescent antibody test for the diagnosis of infectious laryngotracheitis in chickens. Avian Dis. 35:389-391. 1991.

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44. Williams, R A, M Bennett, J. M. Bradbury, R. M Gaskell, R C. Jones, and F. T.W. Jordan. Demonstration of sites of latency of infectious laryngotracheitis virus using the polymerase chain reaction. J. Gen Virol. 73:2415-2420.1992. 45. Williams, R. A., C. E. Savage, and R. C. Jones. A comparison of direct electron microscopy, virus isolataion and a DNA amplification method for the detection of avian infectious laryngotracheitis virus in fiels material. Avian Pathol. 23:709-720. 1994. 46. York, J. J., and K. J. Fahey. Diagnosis of infectious laryngotracheitis using a monocloal antibody ELISA. Avian Pathol. 17:173-182. 1988.

Sholz E., Porter RE. & Guo P. 1994. Differential diagnosis of infectious laryngotracheitis from other avian respiratory disease by a simplified PCR procedure. J. Virol. Meth. 50:313-322. 1994 42. Trist, Η. M, S. G. Tyack, M A. Johnson, C. T. Prideaux, and M Sheppard. Comparison of the genomic short regions of a vaccine strain (SA2) and a virulent strain (CSW-1) of infectious laryngotracheitis virus (Gallid Herpesvirus 1). Avian Dis. 40:130-139.1996. 43. Wilks, C. R. and Kogan, V. G. An immunofluorescence diagnostic test for avian infectious laryngotracheitis. Aus. Vet. Journal. 55:385 - 388. 1979. 41.

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22 MAREK’S DISEASE Patricia S. Wakenell and Jagdev M. Sharma

SUMMARY. Marek’s Disease (MD) is a herpesvirus-induced neoplastic disease of chickens. Because MD virus (MDV) is ubiquitous and chickens acquire environmental infection early in life, they must be protected by vaccination in the hatchery. Diagnosis of infection versus diagnosis of disease is critical as most birds are infected without developing clinical disease. The molecular structure of MDV has been extensively examined. The virus has three serotypes, serotypes 1, 2, and 3. Among these, only viruses of serotype 1 have pathogenic potential. MDV isolates can be categorized by serotype-specific monoclonal antibodies. Exposure to MDV results in persistent infection that lasts for the life of the bird. Infected birds develop viremia as well as antibodies against the virus. Although initially the virus causes a lytic infection in lymphoid cells, the virus persists in the birds by establishing a long-term latent infection of lymphocytes. Latently infected cells minimally express viral antigens. Some latently infected cells may undergo transformation and induce tumors. Thus, MD outbreaks are characterized by development of lymphoid tumors in viscera, skin, and nervous system. MD outbreaks may be associated with mortality, immunosuppression, or excessive condemnation of carcasses during processing. Commercial chicken flocks can experience MD outbreaks at any age. A number of vaccines are available to control MD in the field. These vaccines consist of serotype 1 (attenuated), 2, or 3 viruses administered singly or in combination. Herpes virus of turkeys (HVT), a serotype 3 virus once used as a monovalent vaccine against MD, is now usually mixed with serotype 1 and 2 viruses. Most commercial broiler chickens in the US receive MD vaccines by in ovo technology, in which the vaccine is mechanically injected into eggs at embryonation day 18. Agent Identification. Because most commercial chickens with or without MD have circulating MDV, mere virus isolation or detection of antibody is not helpful in establishing a firm diagnosis. History of the flock, vaccination protocols, the nature of lesions, viral load, expression of viral antigens, and the identity of the cells constituting tumors must be carefully considered. A number of virus isolation strategies are available. Virus isolation in cell cultures is preferred. Because MDV is highly cell-associated, viable cells removed from the tissues or tumors of infected chickens are generally co-cultivated with susceptible monolayer cells to isolate the virus. Upon co-cultivation, the latently infected cells transmit the infection to permissive cells and induce herpesvirus-type cytopathology. The specificity of the cytopathic effect can be confirmed by staining the cell culture monolayers by the immunofluorescence test using antiMDV antibodies. A number of molecular techniques have been used to identify viral genome in tissues. These include polymerase chain reaction, dot-blot hybridization, and in situ hybridization. Assessment of MD viral load in tumors by quantitative PCR and expression of meq on tumor cell surfaces are promising techniques for confirmation of MDV as the causative agent. Serologic Detection in the Host. The serologic tests that can identify anti-MDV antibodies include immunofluorescence, immunohistochemistry, enzyme-linked immunosorbent assay, the agar gel preciptin test, and virus neutralization test. paralysis and/or dilation of the crop due to vagus nerve paralysis. Affected nerves appear edematous, grayish and devoid of cross striations. Blindness is associated with lymphoid infiltration of the iris. Microscopically, MD lesions are composed of an infiltration by a heterogenous population of mononuclear cells.

INTRODUCTION

Marek’s disease (MD) is important primarily in chickens, to a much lesser degree in quail, and has been rarely observed in turkeys, pheasants and jungle fowl. Turkeys and other species have limited susceptibility. Attempts to infect mammals with MD virus (MDV) have uniformly failed. The disease is caused by a widespread, highly contagious, cell-associated, oncogenic herpesvirus. In susceptible chickens, exposure to pathogenic MDV may result in a progressive, debilitating disease that may result in high mortality, reduced egg production, and immunosuppression. The disease most commonly occurs in young, sexually immature chickens 2-7 mo old, but can occur at virtually any age beyond 3 wk. Most commercial flocks are routinely vaccinated against MD. Monovalent or multivalent vaccines are used. Although extensive use of vaccines has greatly reduced the incidence of disease outbreaks compared to the pre-vaccination era, MD occasionally occurs in the presence of vaccine use. Many outbreaks in recent years have been caused by highly virulent strains of the virus that emerged from less virulent predecessors. MDV vaccines protect against the disease but not against infection with the virus. Upon exposure to MDV, vaccinated or unvaccinated chickens become carriers of the virus and persistently shed MDV into the environment, thus making eradication difficult. MDV is not transmitted vertically. The disease occurs throughout the world and virtually all flocks are exposed to the causative virus.

SAMPLE COLLECTION

Since MDV is ubiquitous among chickens, mere demonstration of the virus or antibody cannot be considered pathognomonic for the cause of death or of an eportnic. The disease itself is not ubiquitous and differentiation from other agents causing lymphoid tumors can be problematic (19, 46, 51). For a positive diagnosis of MD in a flock, one must consider the history of the flock and the presence of characteristic gross and microscopic lesions in clinically sick and dead chickens (52). If virus isolation is attempted from commercial chickens raised under field conditions, several MDV types may be isolated. Chickens acquire infection with these viruses through live vaccines or through environmental exposure. The isolated viruses may include herpesvirus of turkeys (HVT, designated serotype 3 MDV) or serotype 1 or 2 MDV. Optimum sources are the same for isolation of HVT as for isolation of MDV. Most suitable for virus isolation are blood and cellular suspensions of spleen or tumor tissue. Isolation from Blood Blood is preferred for virus isolations when euthanasia of the bird is not permitted. In susceptible chickens inoculated experimentally at 1 day of age, viremia peaks at about 4 wk, and most chickens remain persistently viremic for life (52). Infectivity in the blood is associated with the leukocyte (lymphocyte) fraction. The test sample consists of 0.2 ml of whole blood or 2 x 106 buffy-coat cells suspended in cell-culture medium. The buffy-coat cells are the preferred sample.

CLINICAL DISEASE

Clinical signs occur in chickens affected with MD but are of little help in establishing a diagnosis. The disease is characterized by proliferation of lymphoid cells in various tissues and organs, including peripheral nerves. Birds with visceral tumors are depressed and often cachectic prior to death. Birds with lymphoid infiltration of peripheral nerves may demonstrate asymmetric partial 99

Patricia S. Wakenell and Jagdev M Sharma

Isolation from Tumors Solid tumor, spleen, kidney, or other tissues are removed aseptically, minced with scissors, washed several times with phosphate-buffered saline (PBS) until reasonably clear of erythrocytes, and then trypsinized once or twice to obtain single-cell suspensions. The cells are pelleted by centrifugation, resuspended in either PBS or cell-culture medium, and adjusted to give 2 x 106 cells per 0.2 ml. For best results, the cellular preparations should be used immediately after they are processed. If storage is required, the following method is recommended: leukocytes or tissue cells suspended at 2 x 107 cells/ml in cell-culture medium containing ΙΟ­ Ι 5% dimethyl sulfoxide (DMSO) and 15-25% bovine fetal serum are dispensed in 1 ml amounts in glass or plastic cryovials. The ampules are heavily insulated with paper towels and/or placed in a Styrofoam container and stored overnight at -70 C. Ampules are then rapidly, to avoid any thawing, transferred to storage in liquid nitrogen. Cellular suspensions prepared as above can be held at -196 C indefinitely without loss of viability. At the time of use, the ampules are removed from the liquid nitrogen and quickly (< 2 min) thawed in tepid water. The ampules are placed on wet ice and used immediately. For shipping, ampules must be transferred to either liquid nitrogen dewers (preferred) or dry ice without any thawing at any stage of shipment or packaging.

Samples for Antibody Either serum or plasma can be used. Samples are stored and shipped at -20 C. Sera can be shipped at room temperature if a preservative (e.g. merthiolate, benzalkonium chloride or chloroform) is added to prevent bacterial growth.

Isolation of Cell-Free Virus This procedure is not routine. However, for special purposes (e.g., the serum-neutralization test or cloning the virus by plaque purification in cell cultures), cell-free MDV can be extracted from feather tips or skin of infected chickens. For skin preparations, feathers are clipped off at the surface of major feather tracts, skin strips are removed, minced with scissors, and suspended in sucrosephosphate-glutamine albumin (SGPA) buffer containing sodium ethylenediamintetraacetate (EDTA). A 1:5 or 1:10 (w/v) suspension of skin strips in SGPA-EDTA buffer is homogenized for 3-5 min and then sonicated for 2 min (four bursts of 30 sec each) with an ultrasonic oscillator with the needle probe set at an intensity of 70 on the meter. The resulting suspension is centrifuged at 650 x g, and the supernatant is saved as a source of cell-free virus (9). This preparation can be filtered through a 0.45 mm filter pretreated with bovine fetal serum. Cell-free virus can also be isolated from feather tips. Feathers are pulled from all major feather tracts, and 3-5 mm parts of the tips are cut with scissors, diluted 1:5 (w/v) with SPGA-EDTA buffer, and sonicated for 2-3 min. The suspension is cleared by centrifugation and tested for cell-free virus either with or without filtration through a 0.45 mm filter. Cell-free virus preparations should be stored at -70 C in glass or plastic cryovials. For shipment, ampules should be packed in dry ice.

Cell Culture Cell-Associated Virus. Although cell-associated MDV propagates in most avian cell cultures tested, chicken kidney (CK) cells, chicken embryo fibroblasts (CEFs) and duck embryo fibroblasts (DEFs) are the most commonly used. Most classical isolates of serotype 1 MDV replicate poorly or become inconsistent in biologic characteristics when cultivated in CEFs (35). Thus CEFs are generally considered unsuitable for primary virus isolation or propagation for challenge of these viruses. Serotype 2 and 3 viruses grow readily in CEF. Serotype 1 viruses, passed first in susceptible cells, also adapt to grow in CEF. A cellular suspension (0.2 ml containing 2 x 106 cells) is inoculated onto each monolayer culture of 24-hr primary CK cells or secondary CEF or DEF, grown in 15 x 60 mm plastic tissue culture plates. CEF and DEF cultures can be maintained in flasks or roller bottles as well. However, CK maintenance for 5-6 days in flasks or roller bottles is difficult. The cultures are incubated at 37-38 C in a humidified atmosphere containing 3-5% CO2. The culture medium is first renewed 24 hr post-inoculation (PI) and then on alternate days. For serotype 1, foci of CPEs, usually termed plaques, are counted under a light microscope 4-8 days PI in CK cells and 10-14 days PI in DEFs. Direct culture of kidney cells from infected chickens can be done (53). However, kidney cells from older birds may not culture well due to cellular age. Kidney cells from older birds can be used to infect CK monolayers as described. Cell-Free Virus. Cell-free serotype 1 MDV propagates best in primary CK cell cultures. DEF and CEF are less susceptible than are CK cells. Virus is diluted in SPGA-EDTA buffer (Table 1). The SPGAEDTA buffer enhances the virus titer several fold over that with a standard diluent such as PBS. Primary CK cultures, 24-48 hr old, are drained and inoculated with the virus dilutions, two cultures per dilution using 0.2 ml of inoculum per culture. After 30 min of adsorption at 37 C, the medium is added to the cultures. Thereafter, the medium is changed at 48 hr intervals. Because the virus suspension contains EDTA, a chelating agent, the inoculated cells tend to detach from the dish during adsorption. Upon addition of fresh medium, however, the cells resettle and estabfish monolayers. Plaques are counted under the light microscope 6-8 days PI, and the number of plaque-forming units (PFU) per milliliter of virus is calculated.

PREFERRED CULTURE MEDIA AND SUBSTRATES

The best and most widely used substrate for virus isolation is permissive cell cultures. In cell cultures, MDV produces cytopathic effects (CPEs) typical of herpesviruses. Although serotypes 1, 2, and 3 produce herpesvirus-type CPEs, it is possible to differentiate between serotypes by staining CPE-bearing cell cultures with serotype-specific monoclonal antibodies using an indirect immunofluorescence test (26). An experienced eye may also be able to differentiate the morphogenic characteristics of the plaques induced by each serotype. Serial passages of individual isolated plaques picked from an agar-overlaid culture containing a mixture of viruses may be used to prepare stocks of single serotypes. Virus can be isolated by inoculating test materials into embryonating chicken eggs. Isolation of virus by inoculation of susceptible chickens is cumbersome and not commonly done, although the identity of virus isolated in cell cultures or embryos can be confirmed by reproducing typical disease in chickens.

Table 22.1: Sucrose-phosphate-glutamine-albumin buffer containing sodium ethylene diaminetetraacetic acid (EDTA) for extraction and titration of cell-free Marek’s disease virus. The buffer is sterilized by filtration and has a pH of about 6.5 Reagents Concentration Weight Sucrose 0.2180 M 7.462 g Monopotassium phosphate 0.052 g 0.0038 M Dipotassium phosphate 0.0072M 0.125 g L-Monosodium glutamate 0.0049 M 0.083 g Bovine albumin powder 1.0% 1.000 g EDTA 0.2% 0.200 g Distilled H20 qs adA 100 ml A qs ad = sufficient quantity to make

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Chapter 22

Marek’s Disease

The molecular structure of MDV has been examined extensively. The viral DNA for molecular analysis can be obtained by extraction total cell DNA from cell cultures exhibiting extensive viral CPEi, feather tips, feather follicle epithelium (FFE), lymphoid tissues aid brain. Separating virus-specific from cellular DNA can be difficult because the densities of viral and cellular DNA are similar. Many methods are used but pulse field electrophoresis is recommended (48). Cloning of MDV in bacterial artificial chromosomes (BAC) can generate larger amounts of MDV DNA (36). It has been suggested that viral DNA is integrated in the host DNA at multiple sites in chromosomes and the integration pattern varies in different cell lines derived from in vivo MDV-induced tumors (16,24). The complete sequence has been obtained for the 3 serotypes (1,23,41,44). The genome consists of long and short unique regions flanked by terminal repeats (52). Endonuclease digestion patterns differ substantially between the three serotypes (41). Cross hybridization of DNA among the three serotypes under less stringent conditions has shown that these viruses have a collinear relationship. The three types of viruses share significant homology at the DNA level and the structure of some of die dominant genes such as gB, gC, gD, and gH is quite similar within all serotypes (1,23,41,44). There are few structural differences between serotype 1 pathotypes (41). All three serotypes of MDV can cause latent infection in cells but only unattenuated serotype 1 MDV has oncogenic potential. Latently infected cells have limited or no production of virus and have fewer than 5 copies of the viral genome per cell. This number may increase to 15 copies when the cell undergoes neoplastic transformation (20). Viral DNA of transformed cells is highly methylated. Although the molecular basis of oncogenic transformation by MDV is not clear, the gene meq, containing a basic leucine zipper resembling the jun/fos oncogene family, is a strong candidate (25). Some of the transformed cells express meq which can assist in the diagnosis of MDV-induced lymphomas (19,24). The region containing 132 base pair tandem repeats flanks the unique long portion of the virus genome. This region is important because the number of copies of 132 base pair repeats increases following cell culture attenuation of serotype 1 viruses, and thus can be used as a marker for reduced oncogenicity (40). Identification by Inoculation in Cell Cultures, Chickens, or Embryonating Eggs. MDV may be identified by inoculating the test sample in avian cultures (preferably CK cells or DEFs), 1-dayold susceptible chicks, or 9 to 12 day-old embryonating chicken eggs. Positive samples will induce herpesvirus-type CPE in cell cultures, MD in chickens, and pocks on the CAM of eggs. Nonpathogenic isolates of MDV, such as serotypes 2 or attenuated serotype 1, will not induce clinical MD in chickens, although the chickens will develop specific antibodies. In cell cultures, MDV produces CPE characteristic of herpesviruses. The CPEs are sensitive to inhibitors of DNA synthesis such as 5’-iodo-2’-deoxyuridine (IDU). Within 3-5 days of inoculation of monolayer cell cultures with MDV, plaques of CPE appear. These consist of collections of rounded refractile cells. With continued incubation, each plaque grows by including additional refractile cells at its periphery. In Giemsa stained preparations, the refractile cells appear as small or large polykaryocytes. Intranuclear inclusions (Cowdry type A) are readily seen in the areas of CPEs. MDV adapted to grow in cell cultures by serial passage produces large plaques that often develop clear areas (holes) in the center. The morphology of CPE may also vary with the type of cell culture. For instance, in DEF cultures, a focus of CPEs consist of round as well as fusiform cells. In CK cell cultures, fusiform cells are rare. The serotype of the CPE inducing virus can be confirmed by staining infected cultures by the IF test using serotype specific monoclonal antibodies.

Chicken Embryo Inoculation Cell-Associated Virus. Primary virus isolation can be done also by inoculating test material into the yolk sac of 4-day-old embiyonating chicken eggs (45) or intravenously in 10 to 11-dayold embryos. This procedure is rarely used. Virus isolation in cell cultures is preferred. Inoculated and uninoculated control eggs are incubated at 37 C and candled daily. Embryos dying within 24 hr of inoculation are discarded, and those dying thereafter are examined for pocks on the chorioallantoic membrane (CAM). When the embryos are 18 days old, the test is ended by chilling the eggs overnight at 4 C. Positive isolations are characterized by the appearance of pocks on the CAM. One disadvantage of primary virus isolation in embryos is that lymphoid cells present in virusfree inocula may also occasionally produce pocks on the CAM (5). Futhermore, adaptation of MDV to cell cultures by serial passages reduces the pock response by embryos (38). Cell-Free Virus. Inoculum is deposited in the CAM of 9 to 12day-old embryonating eggs, and the CAM is examined for pocks 7 days PI. Cell-free virus can be inoculated intravenously into 10 to 11-day-old embryos (0.1 ml per embryo), and the CAM is examined for pocks 5 days later.

Chicken Inoculation A 0.2 ml test inoculum containing cell-associated or cell-free virus is injected intra-abdominally in 1 to 5-day-old chicks of a susceptible genetic line lacking maternal antibodies to MDV or HVT. The chickens are raised in an isolation environment until they are 6-8 wk old (50, 53). Because MDV is ubiquitous and highly contagious, strict isolation must be maintained to prevent exposure to extraneous virus. Chickens that die before the end of the study should be necropsied, and if gross lesions are absent, sections of nerves (vagus nerves and brachial and sciatic plexuses and nerves), gonads, spleen, kidney, liver and heart should be fixed in 10% neutral buffered formalin and hematoxylin and eosin stained sections examined for histologic changes (30). At the end of the study, survivors are bled for serum and examined for gross and microscopic pathology. Polymerase chain reaction (PCR) can also be performed on tumor and spleen (2,3,4,21,40,55). Antibody can be tested by the agar-gel precipitin (AGP) test, immunofluorescent (IF) test, or ELISA. Groups of chickens infected with the virus develop lesions or antibody or both. Certain highly virulent strains may cause a high incidence of mortality within the first week of infection (49,50). This early mortality is associated with extreme lymphoid cell depletion.

AGENT IDENTIFICATION Physiochemical and Molecular Characteristics of MDV. MDV is ether-sensitive, contains DNA, and belongs to the herpesvirus group (52). The nuclear capsid is 90-100 nm in diameter and contains 162 capsomeres arranged in an icosahedral symmetry. The DNA of MDV is similar for all 3 serotypes and is a buoyant density of 1.705 g/ml. The guanine plus cytosine content is different amongst the 3 serotypes ranging from 43.9-53.6% for serotypes 1 and 2 and 47.6% for serotype 3.MDV remains highly cellassociated in vitro, and transfer of infection from one culture to another depends on the use of viable infected cells. Destruction of cell viability generally results in the loss of infectivity. However, small amounts of cell-free virus can be extracted from cell cultures by SPGA buffer (9). Cell-free MDV obtained from extracts of skin or feather pulp retains its titer for several months at -60 C but not at -20 C. Virus is inactivated in 2 wk at 4 C, in 4 days at 22-25 C, in 18 hrs at 37 C, in 30 min at 56 C, and in 10 min at 60 C. The virus can withstand at least four freeze-thaw cycles and 10 min of ultrasonic vibration. A pH of lower than 4 or higher than 10 readily inactivates the virus. 101

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Virus Detection by Molecular Techniques. These procedures are not routinely used for diagnosis of MD but have recently become popular research tools. Polymerase chain reaction (PCR) technology can be used to identify viral nucleic acid in productively and latently infected cells as well as in cells from lymphomas. The most commonly used primers amplify the 132 base pair repeat sequences (4,21,40,55), although primers to amplify other regions of the genome may also be used (4,19,21). The 132 base pair repeat primers can detect virulent or attenuated serotype 1 MDV. Quantitative PCR has been used to detect MDV in latently infected birds (6,33). Real-time PCR has been used to quantify MDV in tumors (19). Briefly, DNA was extracted from tumors using a Puragene DNA Isolation kit (Gentra System, Minneapolis, MN). Samples were amplified using primers for glycoprotein B (gB) and glyceraldehydes-3-phosphate dehydrogenase (GAPDH). The GeneAmp 5700 (Applied Biosystems, Foster City, CA) was used to amplify the samples in a 25 μΐ PCR reaction containing 50 ng DNA, 0.2pm of each primer and SYBR green master mix (Applied Biosystem, Warrington, UK). The reaction was cycled 40 times. The threshold cycle was determined for each PCR reaction by establishing a fixed threshold. The relative number of copies of gB was compared to that of GAPDH, the reference control (19). High loads of MDV DNA in tumors were considered diagnostic of MDV as the causative agent.

Samples containing oncogenic serotype 1 MDV and HVT may not induce MD in chickens because of the protective effect of HVT. To identify MDV in such samples, inoculated chickens should be maintained in direct contact with uninoculated hatchmates. If oncogenic serotype 1 MDV is present, the contact exposed chickens will not be protected against MD by this virus. If the test samples contain serotype 1 and serotype 2 MDV, neither the inoculated chickens nor the contact exposed chickens will develop MD, although both groups of chickens will develop anti-MDV antibody and persistent viremia with both serotypes. Several pathotypes of serotype 1 MDV have been isolated from chicken flocks. Periodically, highly virulent isolates of serotype 1 virus have emerged that have increasingly resisted protection by conventional vaccines. Thus, serotype 1 MDV isolates recovered from field flocks may vary greatly in virulence. The isolates may be mildly virulent (mMDV), virulent (vMDV), very virulent (wMDV), or very very virulent (w+MDV) (49). These pathotypes can be differentiated by chick inoculation (50,51) although the process is very tedious and should be attempted only by well equipped laboratories proficient in tissue culture techniques and having good animal isolation facilities. Briefly, the virus is first isolated from the test sample in cell cultures. The isolated virus is cloned into a population that contains pure serotype 1 virus without detectable contamination with other serotypes of MDV or other common viruses that may have been present in the test sample. It is imperative that the test sample is free of extraneous viruses that might influence the behavior of the test sample (50,51). The pathotype of the isolated virus is determined by inoculation in chickens that are either unvaccinated or have been immunized with monovalent HVT, bivalent HVT + SB-1 (serotype 2 MDV) or Rispens (attenuated serotype 1) vaccines. All pathotypes cause clinical MD in unvaccinated chickens with the exception of certain highly inbred resistant lines. mMDV is protected by HVT, w MDV is protected by HVT/SB-1 and w+MDV is fairly well protected by Rispens. Direct Demonstration of Viral Antigens in Tissues. Viral antigens can be detected only in tissues that are productively infected with MDV. The FA test using monoclonal or polyclonal ant-MDV antibodies may be used. Fresh tissues are frozen at -70 C or -196 C and cut at 6-8gm thickness in a freezing microtome. At the time of staining, tissue sections are thawed, mounted on glass microscope slides, and stained by the direct or indirect FA test. Viral antigen in the feather follicles may also be detected by reacting feather pulp with anti-MDV polyclonal antibody in an AGP test. The immunohistochemistry (IHC) test using an avidin-biotinperoxidase complex (Vecta-stain ABC kit, Vector Laboratories, Burlingame, CA) has been done to detect meq, CD30, MATSA, p53, pp38 and methyl-3-cytosine in tumors (19). Expression of meq was considered specific for MD lymphomas. Briefly, the IHC staining of meq and CD30 were amplified with the tyramide signal reaction using the TSA Biotin System Kit (PerkinElmer Life Science, Boston, MA). The monoclonal antibody (mAb) for meq (27) was used at a dilution of 1:1000, mAb for pp38 (Hl9.47, 14) was used at a 1:3200 dilution, mAb for AV37, specific for chicken CD30 (7) was used at a dilution of 1:25, the mAb for MATSA (14B367 Lee, unpublished data) was used at 1:2000, the mAb for methyl-3-cytosine (WWR International Oncogene Research Product, San Diego, CA) was used at 1:25 and the mAb for p53 is specific for mouse p53 but cross reacts with chicken p53 (Pab 240, 18) was used at a dilution of 1:25. Molecular procedures can be used to detect the presence of viral DNA in tissues or cells in which viral antigens are not expressed. Dot-blot and in situ hybridization have been used to detect viral antigen in feather tip extracts and to localize viral antigens in tissues, respectively (15,17,22).

Strain Variability Numerous isolates of MDV have been described. Serologically, MDV isolates can be differentiated into sefbtypes 1 and 2. Serotype 1 contains pathogenic isolates and their attenuated forms, and serotype 2 contains apathogenic isolates. Serotype 1 isolates have been differentiated primarily on the basis of pathogenicity for chickens. Thus, they fall into one of four general categories: mMDV, vMDV, wMDV, or w+MDV. Although isolates of serotypes 1 and 2 cross-react, isolates within each serotype have stronger serologic cross-reactions than the isolates between the two serotypes. Highly virulent isolates of MDV (wMDV or w+MDV) are of interest because HVT is not an effective vaccine against these isolates, and bivalent vaccines containing HVT and serotype 2 MDV, certain attenuated serotype 1 vaccines or trivalent vaccines containing HVT along with serotypes 1 and 2 MDV must be used to protect flocks. Diagnosis of infection with wMDV in a flock is a complicated process that has been detailed previously (50). Using real-time reverse transcriptase PCR (54) and/or quantitative PCR to determine viral load (19) show promise for differentiation of pathotypes. Although expansions of the 132 base pair repeat have indicted attenuation, these are not related to virulence (42). Mutations in genes such as meq (10) and the terminal and internal repeat long regions of MDV may also contribute to differences in virulence (43). Characteristics of Vaccine Strains Although HVT was once the most commonly used vaccine against MD, many flocks now receive bivalent or trivalent vaccines. Bivalent vaccines contain HVT and a serotype 2 MDV (e.g., SB-1), whereas trivalent vaccines may contain HVT, attenuated serotype 1 MDV (e.g., CV1988), and serotype 2 MDV (e.g., SB-1). In some parts of the world, serotype 1 MDV is used alone as an effective vaccine against MD. Recombinant MD vaccines have been developed that are quite similar to HVT/SB-1 in protecting chickens against virulent MDV. The recombinants have been constructed by inserting immunogenic genes of MDV, particularly gB, in fowlpox virus or HVT (28,34). Recombinant vaccines containing immunogenic genes from multiple disease agents such as MDV and Newcastle disease virus have also been developed (32). Upon vaccination, chickens remain asymptomatic, although they develop persistent viremia with all viruses present in the vaccine.

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the medium, whereas the other lines are minor, produced by antigens intimately associated with infected cells.

Vaccinated chickens become resistant to tumor formation by virulent MDV but not to infection with MDV. Thus, the vaccinated chickens become persistently viremic with the vaccine virus(es) as well as field MDV and can shed virulent MDV, although at a reduced rate. The best way to ascertain that the flock has been properly vaccinated against MD is to isolate the vaccine virus(es) from vaccinated chickens, HVT replicates readily in avian cell cultures (8), although CPEs are slightly different from that of MDV. In contrast to MDV plaques, which develop slowly and consist of rounded clusters rarely extending more than 0.5-1 mm in diameter, the plaques produced by HVT in CK cells appear early (in 2-3 days) and consist of large syncytia surrounding a clear central area. HVT plaques grow rapidly, reaching a diameter of 1.5-3 mm in about 10 days. HVT and MDV plaques can be distinguished in several avian cells cultures, but most clearly in CK cell cultures. The identity of the plaques induced by the two viruses may be confirmed by staining CPE-monoclonal antibodies in an FA test. Serotypes 1 and 2 antibody will react only with MDV plaques and serotype 3 antibody reacts with HVT plaques. Most MD vaccines consist of a suspension of viable, virus-infected CEF (wet vaccine), although lyophilized, monovalent cell-free HVT vaccine (diy vaccine) is also available. Serotypes 1 or 2 MDV are not available in the lyophilized form. Wet vaccine is supplied in frozen, sealed glass ampules. The accompanying handling instructions should be followed carefully. The vaccine should be thawed just before used and immediately diluted to the proper dose level. Concentrations of DMSO used to protect cells during freezing are highly toxic to cells at room temperature (25-30 C).

Fluorescent Antibody Test The FA test can be used to detect antibodies if known antigen is available (indirect FA) or - the reverse - to detect antigen if know® antibody is available (direct FA). Techniques for both direct and indirect FA tests for the avian system have been reviewed (31). Far the direct test, globulin extracted from serum of chickens exposed to MDV is labeled with fluorescein isothiocyanate. For the indirect test, labeled anti-chicken gamma globulin is available from several commercial sources. Mouse monoclonal anti-MDV antibodies have been developed that may be used in the indirect FA test (26). The antigen is prepared in monolayer cultures of CK cells grown on glass coverslips. Confluent monolayers are inoculated with enough MDV to give numerous separated plaques. When the earliest distinct plaques appear (before CPE become confluent), the coverslips are washed once with PBS and then fixed for 5 min in acetone at 4 C. Fixed coverslips can be stored at -20 C and used within several weeks. Enzyme-Linked Immunosorbent Assay In this test, wells of a 96-well microtiter plate are coated with MDV-infected CEFs or DEFs (11). Monolayer cells cultured in 100-mm plates are infected with a massive dose of cell-associated MDV. When more than 75% cells show CPEs, the infected cells are trypsinized and suspended in PBS and used to coat the wells of the microtiter plate. Alternatively, the cells may be suspended in tissue­ culture medium containing 10% calf serum and 10% DMSO and stored in liquid nitrogen for later use. At the time of use, the frozen cells are thawed and washed twice with PBS to remove the DMSO before coating the wells. Each microtiter well receives 5 x 104 infected cells (containing about 1.7 x 103 PFU) in 0.1 ml PBS. After the plates are low-speed centrifuged (1000 rpm) for 10 min, the supernatant is discarded and the cells are allowed to dry at room temperature. Antigen-coated plates may be stored at 4 C or -20 C for at least 3 mo without loss of reactivity. The stored plates should be washed three times with PBS containing 0.1% Tween 80 before use. Duplicate samples of 0.1 ml of test sera and known MD antibody­ positive and -negative sera diluted in 3% bovine serum albumin are placed in the wells of the microtiter plates. After 1 hr at 37 C, the wells are washed with PBS and 0.1 ml of a 1:3200 dilution of affinity-purified goat anti-chicken immunoglobulin G (IgG)peroxidase conjugate (Synbiotics, San Diego, CA) is added to each well. After 1 hr at 37 C, the wells are washed four times and to each well is added 0.1 ml of substrate containing one volume of 0.05% H2O2 and nine volumes of purified 5-aminosalicylic acid (1 mg per ml in 0.02 M phosphate buffer [pH6.0]). After 30 min at room temperature, absorbance is read at 490 nm wavelength using any commercially available ELISA reader. Sera showing an absorbance reading of at least three times that of known antibody-negative sera are considered positive. Cheng et al. (11) considered a serum positive if the absorbance at a 1:400 dilution was 0.20 units or higher. The titer of antibody was the reciprocal of the serum dilution with an absorbance of 0.20 units.

SEROLOGIC DETECTION IN THE HOST

Chickens infected with MDV develop antibody that persists for life. Several serologic tests may be used to quantify serum antibody, however, usefulness in commercial flocks is limited since all flocks have been vaccinated or exposed. Serologic assays are useful for evaluating chickens with no known prior exposure to MDV vaccine or field strains (backyard flocks, international flocks). Agar-Gel Precipitin Test In the AGP test, serum is reacted with MD antigens in an agar medium. Commercial reagents are available. The antigen for this test is prepared by propagating MDV (of low cell-culture-passage level) in CK cell or CEF cultures (12). When CPEs are confluent, the cells are detached from the culture vessel either by scraping the vessel with a rubber policeman or trypsinizing and the cells are suspended in PBS or culture medium without tryptose phosphate broth (tryptose phosphate broth in the antigen may produce nonspecific precipitin bands) at a concentration of about 1 x 107 cells/ml. This suspension is then freeze-thawed three times and used as antigen. Feather pulp extracted by squeezing the tips of feathers plucked from major feather tracts of infected chickens also has been used as antigen. For optimum precipitation, a 1% suspension of Noble agar is made in NaK phosphate buffer (pH 7.4) containing 8% NaCl. About 3 ml of suspension is poured onto a glass microscope slide pretreated with a film of 1:5 dilution of agar suspension. When the agar solidifies, one central and six peripheral equidistant wells, each 3 mm in diameter and about 3.5 mm apart are cut with a template or other suitable borer. The central well is filled with antigen, and the peripheral wells are filled with serum. To detect nonspecific precipitin bands and enhance weak reactions each well containing test serum should be adjacent to one containing known MD-specific serum. Reagents should be diffused for 72 hr in a moist chamber at either 37 C or at room temperature. Positive test serum should form a line of identity with the line produced by MD-specific serum. Certain sera may produce up to six precipitin lines. The most prominent line, referred to as the A (or gC) line, is produced by the antigen released from infected cells into

Virus Neutralization Test This test is used to detect neutralizing antibody in the serum or plasma of infected chickens. Because the cell-free preparations of MDV prepared from skin or feather-follicle extracts of MDVinfected chickens are generally low in titer, the kinetics of neutralization have not been adequately studied. The test should be conducted with cell-free virus suspension in SPGA-EDTA buffer with a titer of about 103 PFU/ml. One part of twofold or 10-fold serum or plasma dilution and four parts of virus suspension are mixed and incubated for 30 min at 37 C or at room temperature.

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7. Burgess, S. C., P. Kaiser, and T. F. Davison. A monoclonal antibody that recognizes the chicken homologue of CD30, a tumor antigen in Marek’s disease. In: K. A. Schat (Ed). Current Progress in Avian Immunology. Am. Assoc. Avian Path. Kennett Square, PA. pp. 232-232. 2001. 8. Calnek, B. W., C. Garrido, W. Okazaki, and I. V. Patrascu. In vitro methods for assay of turkey herpesvirus. Avian Dis. 16:52-56. 1972. 9. Calnek, B. W., S. B. Hitchner, and Η. K. Adldinger. Lyophilization of cell-free Marek’s disease herpesvirus and a herpesvirus of turkeys. Appl. Microbiol. 20:723-726. 1970. 10. Chang, K. S., K. Ohashi, and M Onuma. Diversity (polymorphism) of the meq gene in the attenuated Marek’s disease virus (MDV) serotype 1 and MDV-transformed cell lines. J. Vet. Med. Sci. 64:1097-1101. 2002. 11. Cheng, Y. Q., L. F. Lee, E. Smith, and R. L. Witter. An enzyme-linked immunosorbent assay for the detection of antibodies to Marek’s disease virus. Avian Dis. 28:900-911. 1984. 12. Chubb, R. C., and A E. Churchill. Precipitating antibodies associated with Marek’s disease. Vet. Rec. 83:4-7. 1968. 13. Crespo, R., P. R. Woolcock, A. M Fadly, C. Hall, and H. L. Shivaprasad. Characterization of T-cell lymphomas associated with an outbreak of reticuloendotheliosis in turkeys. Avian Pathology 31:355-361. 2002. 14. Cui, Z. Z., D. Yan, and L. F. Lee. Marek’s disease virus gene clones encoding virus-specific phosphorylated polypeptides and serological characterization of fusion proteins. Virus Genes 3:309-322. 1990. 15. Davidson, I., M Malkinson, C. Strenger, and Y. Becker. An improved ELISA method using a streptavidin-biotin complex, for detecting Marek’s disease virus in feather tips of infected chickens. J. Virol. Methods 14:237241. 1992. 16. Delecluse, H.-J., and W. Hammerschmidt. Status of Marek’s disease virus in established lymphoma cell lines: herpesvirus integration is common. J. Virol. 67:82-92. 1993. 17. Endoh, D., Y. Kon, M Hayashi, T. Morimura, K. O. Cho, T. Iwasaki, and F. Sato. Detection of transcripts of Marek's disease virus serotype 1 ICP4 homologue (MDV ICP4) by in situ hybridization. J. of Vet. Mei Sci. 58:969-975. 1996. 18. Gannon, J. V., R. Greaves, R. Iggo, and D. P. Lane. Activating mutations in p53 produce a common conformational effect. A monoclonal antibody specific for the mutant form. Euro. Mol. Biol. Org. J. 9:1595-1602. 1990. 19. Gimeno, I. M, R. L. Witter, A. M Fadly, and R. F. Silva. Novel criteria for the diagnosis of Marek’s disease virus-induced lymphomas. Avian Pathology 34(4):332-340. 2005. 20. Gimeno, I. M, R. L. Witter, and A. Miles. Marek’s disease (CDRom) Slide set. Am. Assoc. Avian Path. Athens, GA. 2004. 21. Handenberg, K. J., O. L. Nielsen, and P. H. Jorgensen. The use of serotype 1- and serotype 3-specific polymerase chain reaction for the detection of Marek’s disease virus in chickens. Avian Pathology 30:243249. 2001. 22. Holland, M S., C. D. Mackenzie, R. W. Bull, and R. F. Silva. A comparative study of histological conditions suitable for both immunofluorescence and in situ hybridization in the detection of herpesvirus and its antigens in chicken tissues. J. Histochem. Cytochem. 44:259-265. 1996. 23. Izumiya, Y, Η. K. Jang, M Ono, and T. Mikami. A complete genomic DNA sequence of Marek’s disease virus type 2, strain HPRS24. In: K. Hirai (Ed.). Current Topics in Microbiology and Immunology. Springer-Verlag, Berlin. 255:191-222. 2001. 24. Jarosinski, K. W., B. K. Tischer, S. Trapp, andN. Osterrieder. Marek’s disease virus: Lytic replication, oncogenesis and control. Expert. Rev. Vaccines 5(6):761-772. 2006. 25. Kung, H. J., L. Xia, P. Brunovskis, D. Li, J. L. Liu, and L. F. Lee. Meq: An MDV-specific bZIP transactivator with transforming properties. In: K. Hirai (Ed.). Current Topics in Microbiology and Immunology. SpringerVerlag, Berlin. 255:245-260. 2001. 26. Lee, L. F., X. Liu, and R. L. Witter. Monoclonal antibodies with specificity for three different serotypes of Marek’s disease viruses in chickens. J. Immunol. 130:1003-1006. 1983. 27. Liu, J. L., L. F. Lee, and H. J. Kung. Biological properties of the Marek’s disease latent protein MEQ: Subcellular localization and transforming potential. In: R. F. Silva, Η. H. Cheng, P. M Coussens, L. F. Lee, and L. F. Velicer (Eds.). Current Research on Marek’s Disease. Am. Assoc. Avian Path. Kennett Square, PA pp. 271-277. 1996. 28. Nazerian, K., L. F. Lee, N. Yanagida, and R. Ogawa. Protection against Marek’s disease by fowlpox virus recombinant expressing the glycoprotein B of Marek’s disease virus. J. Virol. 66:1409-1413. 1992.

Known positive and negative serum controls should be included in the test. Each of duplicate cultures of 24-hr-old primary CK cells is then inoculated with 0.2 ml of the serum-virus mixture and absorbed for 30 min at 37 C and fresh medium is then added to the cultures. Thereafter, medium is changed on alternate days. Plaques are counted 6-8 days after inoculation. The titer of the serum is the reciprocal of the serum dilution that causes at least 50% reduction in virus titer (obtained with negative serum control).

Vaccine Delivery. Within the last decade, a dramatic change has occurred in the method of MD vaccine delivery in commercial chicken flocks. Previously, MD vaccines were administered subcutaneously in newly hatched chickens in the hatchery. Most of the major hatcheries in the U.S.A. now use in ovo technology to vaccinate chickens against MD (37,39). In this technology, the vaccines are injected in eggs at 17-18 days of embryonation using multiple-head injection machines. Site of vaccine delivery is critical for MDV vaccine efficacy (47). This method has substantially reduced the labor cost associated with posthatch vaccination. DIFFERENTIATION FROM CLOSELY RELATED AGENTS Clinical MD is often confused with lymphoid leukosis. The main source of confusion is the similarity of the gross visceral tumors produced by the two diseases. Detection of cellular antigens with monoclonal antibodies may be used to distinguish tumor cells of MD and lymphoid leukosis (13,19,29,51). Because reticuloendotheliosis virus may cause nerve enlargement or lymphomas under experimental conditions, it also may be confused with MD. Firm diagnosis depends on flock history, distribution and histologic appearance of lesions, identity of cells constituting tumors, identification of viral antigens in tumor cells, and isolation of the etiologic agent (see Chapter 35 on oncornaviruses). Viral antigens may be identified by microscopy techniques using immunofluorescence or immunohistochemhical staining or by molecular assays such as PCR to detect viral nucleic acid. Another common avian herpesvirus that should differentiate from MDV is infectious laryngotracheitis virus (ILTV). Clinically, ILTV causes a respiratory disease in chickens, and the lesions are more or less restricted to the upper respiratory tract, without involvement of the nervous system. ILTV produces pocks that are indistinguishable from those produced by MDV on the CAM of embryonated eggs. In avian cell cultures, ILTV becomes cell-free and grows much faster than MDV. The CPE of ILTV becomes detectable 8-10 hr after inoculation and consists of rapidly progressing syncytia formation. Plaques induced by ILTV and MDV may be differentiated by staining with virus-specific antibodies. REFERENCES

1. Afonso, C. L., E. R. Tulman, Z. Liu, L. Zsak, D. L. Rock, and G. F. Kutish. The genome of turkey herpesvirus. J. Virol. 75:971-978. 2001. 2. Baigent, S., V. Nair, and R. Currie. Real-time quantitative PCR for Marek’s disease vaccine virus in feather samples: applications and opportunities. Dev. Biol. (Basel). 126:271-81. 2006. 3. Baigent, S. J., L. J. Pelherbridge, K. Howes, L. P. Smith, R. J. Currie, and V. K. Nair. Absolute quantification of Marek’s disease virus genome copy number in chicken feather and lymphocyte samples using real-time PCR. J. Virol. Methods 123(l):53-64. 2005. 4. Becker, Y., Y. Asher, E. Tabor, I. Davidson, M Malkinson, and Y. Weisman. Polymerase chain reaction for differentiation between pathogenic and nonpathogenic serotype 1 Marek’s disease virus (MDV) and vaccine viruses of MDV serotypes 2 and 3. J. Virol. Methods 40:307-322. 1992. 5. Biggs, P. M, and B. S. Milne. Use of the embryonating egg in studies on Marek’s disease. Am. J. Vet. Res. 32:1795-1809. 1971. 6. Bumstead, N. J., Silliboume, M. Rennie, N. Ross, and F. Davison. Quantification of Marek’s disease virus in chicken lymphocytes using the polymerase chain reaction with fluorescence detection. J. Virol. Methods 65:75-81. 1997.

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29. Neumann, U., and R. L. Witter. Differential diagnosis of lymphoid leukosis and Marek’s disease by tumor associated criteria. I. Studies on experimentally infected chickens. Avian Dis. 23:417-425. 1979. 30. Payne, L. N., and P. M Biggs. Studies on Marek’s disease. Π. Pathogenesis. J. Natl. Cancer Inst. 39:281-302. 1967. 31. Purchase, H. G. Fluorescent-antibody techniques in avian research. Avian Dis. 17:213-226. 1973. 32. Reddy, S. K., J. M Sharma, J. Ahmad, D. N. Reddy, J. K. McMillen, S. M Cook, M A. Wild, and R. D. Schwartz. Protective efficacy of recombinant herpesvirus of turkeys as an in ovo vaccine against Newcastle and Marek’s disease in specific-pathogen-free chickens. Vaccine 14:469477. 1996. 33. Reddy, S. M, R. L. Witter, and I. M Gimeno. Development of a quantitative-competitive polymerase chain reaction assay for serotype 1 Marek’s disease virus. Avian Dis. 44:770-775. 2000. 34. Ross, L. J. N., Μ M Binns, P. Tyers, J. Pastorek, V. Zelnik, and S. Scott. Construction and properties of turkey herpesvirus recombinant expressing the Marek’s disease virus homolog of glycoprotein B of herpes simplex virus. J. Gen. Virol. 74:371-377. 1993. 35. Schat, K. A. Isolation of Marek’s disease virus: Revisited. Avian Pathology 34(2):91-95. 2005. 36. Schumacher, D. Β., Β. K. Tischer, W. Fuchs, and N. Osterrieder. Reconstruction of Marek’s disease virus serotype I (MDV-1) from DNA cloned as a bacterial artificial chromosome and characterization of a glycoprotein B negative MDV-1 mutant. J. Virol. 74:11088-11098. 2000. 37. Sharma, J. M., and B. R. Burmester. Resistance to Marek’s disease virus at hatching in chickens vaccinated as embryos with the turkey herpesvirus. Avian Dis. 26:134-'49. 1982. 38. Sharma, J. Μ, B. D. Coulson, and E. Young. Effect of in vitro adaptation of Marek’s disease virus on pock induction on the chorioallantoic membrane of embryonated chicken eggs. Infect. Immun 13:292-295. 1976. 39. Sharma, J. M, and R. L. Witter. Embryo vaccination against Marek’s disease with serotypes 1, 2, and 3 vaccines administered singly or in combination. Avian Dis. 27:453-463. 1983. 40. Silva, R. F. Differentiation of pathogenic and nonpathogenic serotype 1 Marek’s disease viruses (MDVs) by the polymerase chain reaction amplification of the tandem direct repeats within the MDV genome. Avian Dis. 36:521-528. 1992. 41. Silva, R. F., L. F. Lee, and G. F. Kutish. The genomic structure of Marek’s disease virus. In: K. Hirai (Ed.). Current Topics in Microbiology and Immunology. Springer-Verlag, Berlin. 255:143-158. 2001.

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23 DUCK VIRUS ENTERITIS Peter R. Woolcock

SUMMARY. Duck virus enteritis (DVE) is caused by a herpesvirus of the subfamily Alphaherpesvirinae of the family Herpesviridae. No evidence exists of antigenic variation. Infections have only been reported in ducks, geese, and swans. Birds of all ages are susceptible to infection and mortality may be high, but the severity of clinical signs varies with the species, sex, and age of the infected birds. Lesions seen at necropsy are typical of vascular damage. Agent Identification. Diagnosis of DVE is confirmed by isolation and identification of the virus. Polymerase chain reactions (PCR) have been developed to detect viral DNA in infected tissues. Serologic Detection in the Host. Serologic detection of infection is only of importance in non-commercial waterfowl, and may be used to determine prior exposure of migratory and non-migratory birds to DVE virus.

INTRODUCTION

SAMPLE COLLECTION

Duck virus enteritis (DVE) is an acute, sometimes chronic, contagious viral infection occurring naturally only in ducks, geese, and swans, all of which are members of the family Anatidae of the order Anseriformes. DVE may also be referred to as duck plague, anatid herpes, eendenpest, entenpest, and peste du canard. The disease has been reported in Asia, Europe, and North America. The etiologic agent, a herpesvirus, is a member of the subfamily Alphaherpesvirinae of the family Herpesviridae (16, 17). The infection has not been reported in other avian species, mammals, or humans.

Carcasses or tissues collected for virus isolation from dead ducks should be chilled at 4 C immediately; if they cannot be delivered to a diagnostic laboratoiy within 24 hr, the tissues should be frozen, preferably at -70 C or lower, and stored until they can be shipped on dry ice (freezing at temperatures warmer than -70 C may result in loss of viability of the virus). Primary isolation of the virus is best achieved from samples of liver, spleen, or kidney tissue that have been homogenized in buffered saline and clarified by low speed centrifugation. PREFERRED CULTURE MEDIA AND’SUBSTRATES

CLINICAL DISEASE

Virus isolation may be attempted by inoculating clarified tissue homogenates onto cell cultures, or into ducklings or duck embryos.

In domestic ducks and ducklings, DVE has been reported in birds ranging from 7 days of age to mature breeders. In susceptible flocks, wild or domestic, the first signs are often sudden, high, and persistent mortality, and a significant drop in egg production. In chronically infected, partially immune flocks, only occasional deaths occur. Latent infections are established in recovered birds and these carriers may shed the virus in feces or on the surface of eggs over a period of years (16,18). Clinical signs and gross lesions associated with a DVE outbreak vary not only with the virulence of the strain of virus, but also with the species, age, and sex of the affected waterfowl (1, 17). In breeder ducks, the signs may include photophobia, polydypsia, loss of appetite, ataxia, watery diarrhea, and nasal discharge. Affected birds often have ruffled feathers and soiled vents. Sick birds may maintain an upright stance by using their wings for support, but their overall appearance is one of weakness and depression. In ducklings 2-7 wk of age, mortality may be lower than in older birds and the signs associated with DVE infection include dehydration, loss of weight, a blue coloration of the beaks, and blood-stained vents. At necropsy, little evidence exists of emaciation in adult ducks. Prolapse of the phallus may occur in mature males. The gross lesions are characterized by vascular damage, with tissue hemorrhages and free blood in the body cavities, eruptions or annular hemorrhages (seen as dark red bands around the intestine), and ulcers with diphtheritic plaques on the mucosal surfaces of the digestive tract. Lesions occur in all the lymphoid organs and degenerative changes are apparent in the parenchymatous organs. Collectively, these lesions are pathognomonic for DVE. The pathology and histopathology of DVE in white Pekin ducks (Anas platyrhynchos) has been reviewed (1, 17).

Cell Cultures Isolation of DVE virus may be made in primary duck embryo fibroblasts (DEFs) (8, 13, 19), or preferably, primary Muscovy DEFs (MDEFs) (8, 13). Cell monolayers grown in medium consisting of Eagle minimal essential medium (EMEM) containing 10% fetal calf serum (FCS), 2 mM glutamine, and 0.17% sodium bicarbonate and gentamicin are washed with serum-free EMEM and then inoculated with the clarified sample homogenate suspected to contain DVE virus. The virus is added to the cell monolayer and incubated for 1 hour at 37 C and then EMEM medium containing 2% FCS, 2 mM glutamine, and 0.17% sodium bicarbonate and gentamicin is added and the cultures are incubated at 37 C in an atmosphere containing 5% CO2 This method of isolation may be modified to a plaque assay by overlaying the monolayer with maintenance medium containing 1% agarose. Primary Muscovy duck embryo liver cells may be more sensitive for virus isolation, (R.E. Gough, pers. comm.). It has been reported (2) that the isolation of DVE virus in MDEF cells is favored by incubation at temperatures between 39.5 and 41.5 C. However, an elevated temperature does not appear to be essential for isolation, which is often carried out at 37 C. More than one passage may be necessary for an isolation. The cytopathic effect is characterized by the appearance of pyknotic, rounded clumped cells that enlarge and become necrotic after 2-4 days. To confirm the presence of DVE virus, cultures should be stained with a fluorescent antibody conjugate using a direct or indirect method specific for DVE virus. Cell monolayers may also be fixed and then stained with hematoxylin and eosin to show syncytial formation, intranuclear inclusions, and marked cytoplasmic granulation. Ducklings When inoculated intramuscularly, 1-day-old susceptible ducklings die within 3-12 days. Muscovy ducklings (Cairina moschata) (Grimaud Farms, Stockton, Calif.) are more susceptible to DVE than are white Pekin ducklings. Both gross and microscopic lesions 106

Chapter 23

typical of infection with DVE should be seen on postmortem examination. Uninoculated ducklings, housed separately, should be maintained as controls. The diagnosis may be confirmed either by vaccinating ducklings against DVE (Cornell University, Duck Research Laboratory, Eastport, Long Island, N.Y.) and challenging them subsequently with the virus isolate, or by staining of tissues with a fluorescent antibody conjugate specific for DVE virus. However, virulent strains of the virus do exist, against which vaccine may be ineffective (12). The susceptibility of Pekin ducklings to field isolates of DVE virus can be highly variable and therefore unreliable as the primary method of isolation. Virus isolation in birds rather than in egg or cell cultures is not always practical, but it may be necessary for a successful demonstration of the virus. In the author's recent experience, even though a herpesvirus could be visualized by negative stain electron microscopy, and the necropsy and histopathologic changes observed in Muscovy ducks were pathognomonic for DVE, isolation of virus was eventually achieved only in Muscovy ducklings and not in MDEF cell cultures. Duck Embryos Primary DVE virus isolations can also be made by inoculation onto the chorioallantoic membrane (CAM) of 9 to 14-day-old embryonating Muscovy duck eggs. The embryos may die, showing characteristic extensive hemorrhages 4-10 days after inoculation. Two to four serial passages of the homogenized CAMs may be necessary before an isolation is made. This method of isolation is not as sensitive as cell cultures or susceptible 1-day-old ducklings. Pekin duck embiyos vary in their susceptibility to strains of DVE virus. Embryonating chicken eggs are not very susceptible to infection with field strains of DVE virus. The virus can, however, be adapted to chicken embryos by serial passaging.

AGENT IDENTIFICATION

Morphology Mature virions are enveloped, have typical herpesvirus morphology, and range in size from 160 to 380 nm. Naked or unenveloped nucleocapsids may be seen by negative stain electron microscopy and are between 100 to 110 nm in diameter. Physicochemical Properties The virus genome consists of DNA and virus particles are sensitive to lipid solvents and are rapidly inactivated at a pH 10.

Duck Virus Enteritis

Biological Properties Infected cells surrounding necrotic foci contain intranuclear inclusion bodies. Based on plaque-reduction tests, all strains of DVE virus currently belong to one serotype. Chicken embryo-adapted DVE virus is used in the preparation of a live attenuated vaccine, which is recommended for use in birds 2 weeks of age or older. Fattening or breeding ducks may be vaccinated subcutaneously or intramuscularly to produce active immunity. The vaccine virus is thought not to spread by contact from vaccinated to unvaccinated ducks because unvaccinated contact birds remain susceptible to infection.

Antigen Detection Virus-neutralization (VN) and immunofluorescence tests are most commonly used to confirm the identity of DVE virus using either embryonating eggs or cell cultures. A plaque assay for DVE virus using duck embryo cell cultures has been described (4). Hyperimmune antiserum prepared in ewes was used in a direct fluorescent antibody test for DVE virus in DEF cells, and was shown to be the next most sensitive assay after isolation in 1 to 9day old ducklings (7). Direct or indirect immunofluorescent tests using antisera to DVE virus prepared in ducks can be used to detect virus in cell cultures and in frozen tissue sections. A reverse passive hemagglutination test for DVE has been described (5) but is reported to be less sensitive than immunofluorescence and plaque assays. An avidin-biotin-peroxidase method of immunoperoxidase staining to detect DVE antigen in formalin-fixed, paraffinembedded sections of liver and spleen from experimentally infected birds has been described (11); this method could have diagnostic potential. Herpesvirus may also be detected in clarified tissue homogenates by negative stain electron microscopy; this is not, however, positive confirmation that the herpesvirus is DVE virus. Molecular identification Detection of DVE virus by polymerase chain reaction (PCR) has been reported (9, 10, 14, 15). Primers have been identified that are able to amplify DNA from DVE virus present in various tissues, including oesophagus, liver and spleen, from an original outbreak and after passage in Muscovy duck embryos The following detailed protocol for the detection of DVE virus was provided by Dr. W. R. Hansen, US Geological Survey, Biological Resources Division, National Wildlife Health Center, 6006 Schroeder Road, Madison, WI 53711, USA. This procedure uses the following commercial items: GeneAmp PCR Reagent Kits containing dNTPs, lOx amplification buffer for hot start PCR, Taq DNA polymerase, Lambda PCR control reagents, and Ampliwax beads (Applied Biosystems), and a 100 base pair molecular size ladder (Invitrogen).

Extraction of viral DNA This DNA extraction procedure can be used on disrupted cell suspensions from duck plague infected cell culture, 10% ground tissue suspensions, or cloacal swab material in transport medium. This method is used to prepare duck plague DNA for the known positive PCR controls. Note: All product transfers in steps i through v are performed inside a biological safety cabinet.

For 10% ground tissue suspension, add 400 μΐ to a 1.5 ml microfuge tube and microfuge at 16,000xg for 5 min. Transfer the supernatant to a new tube and go to step ii. i. For cell culture suspensions and cloacal swab material, add 400 μΐ of the sample, or supernatant from step i above, to a 1.5 ml tube and microfuge at 16,000-20,OOOxg for 45 min, to pellet virus. ii. Discard the supernatant and resuspend the pellet with 200 μΐ of Tris - EDTA (10 mM Tris-HCl, pH 8.0, 1 mM EDTA) buffer. iii. Add 10 μΐ of a 5 pg/μΙ proteinase K solution giving a final concentration of 0.2 pg/μΙ, mix thoroughly, and incubate at 56 C for 1 hour. iv. Add 25 μΐ of 10% sodium dodecyl sulfate solution giving a final SDS concentration of 1%, mix thoroughly, and incubate at 37° C for one hour. v. Add 15 μΐ of 5M NaCl giving a final concentration of 0.3 M and mix thoroughly. vi. Add 300 μΐ of fresh phenol buffered with Tris-HCl, pH 8.0 to the tube, and mix by inverting 50 times. vii. Microfuge tube at 16,OOOxg for 5 min and transfer the top aqueous phase (sample) to a new tube. viii. Repeat the phenol extraction steps vii and viii one more time. 107

Peter R Woolcock

ix. x. xi. xii. xiii. xiv. xv. xvi. xvii.

Add 500 μΐ of ether to the tube, mix thoroughly, and microfuge at 16,000 x g for one min. Discard the top aqueous phase (ether) and repeat the ether extraction step (x) one more time. Heat tube with lid open at 56 C for about 15 min or until the smell of ether is gone. Split tube contents in two and add 2.25 times the sample volume of 100% ethanol to each tube, mix tube contents by inverting tube several times, and leave at room temp (22 C) for 30 min. Microfuge tube at 16,000 x g for 45 min, and discard supernatant. Add 200 μΐ of 70% ethanol to gently wash pellet then microfuge at 16,000 x g for 15 min. Discard the supernatant and diy pellet at 56 C for 30-45 min, with the tube lid open. Resuspend the DNA in 30 μΐ of distilled water that is RNAase and DNAase free. Store sample tube at 4 C until tested (few days) or at -20 C for long term storage.

Polymerase chain reaction i. Lower reaction mixtures for the duck plague PCR and the lambda control are prepared in advance in a biosafety cabinet using the kit manufacturers recommended methods for a hot start PCR. The lower reaction mixture is dispensed into PCR reaction tubes, sealed with Ampliwax at 80 C as recommended by the manufacturer, and stored at 4 C for 1-2 mo. ii. The following are the PCR primers for duck plague DNA-directed DNA polymerase gene: iii. Primer 1 sequence 5’- GAAGGCGGGTATGTAATGTA - 3’ (forward) iv. Primer 2 sequence 5’ - CAAGGCTCTATTCGGTAATG - 3’ (reverse) v. The upper reaction mixture is prepared as a master mix according to the kit manufacturer’s recommendations the day of the test, and distributed to each sample tube including the duck plague and lambda control tubes. vi. Add 10 μΐ of DNA suspension from the stored sample tubes to PCR lower reaction tubes with corresponding labels. vii. Place known duck plague DNA diluted to 1 pg/10 μΐ into one control tube and 10 μΐ of distilled water into the no DNA control tube. Add 10 μΐ of lambda DNA supplied in the kit and 10 μΐ of water to corresponding lambda control tubes. viii. Place all the tubes in a thermal cycler that is programmed as follows: One cycle: Hold 94 C for 2 min. Hold 37 C for 1 min. Hold 72 C for 3 min. 35 cycles: Hold 94 C for 1 min. Hold 55 C for 1 min. Hold 72 C for 2 min. One cycle: Hold 72 C for 7 min. Hold 4 C until stored PCR tubes are stored at 4 C until samples are examined for amplification products.

Electrophoretic analysis of PCR products i. A fresh lx TAE buffer (40 mM Tris acetate, 1 mM EDTA, pH 8.3) is prepared from a lOx stock for agarose preparation and for use in the electrophoresis chamber. ii. A 1% agarose solution is prepared in TAE buffer, heated to dissolve the agarose and, when cool, poured into a gel former with a comb. iii. The solidified gel is placed into the electrophoresis chamber and TAE running buffer is added. iv. PCR test samples, including the duck plague and lambda controls, are mixed 1/10 with Ιμΐ of loading buffer (0.25% (w/v) bromophenol blue, 0.25% (w/v) xylene cyanol, 0.01 M Tris-HCl, pH 8.0, and 50% (v/v) glycerol) and 10 μΐ of each is added to individual wells of the gel. The 100 base pair (bp) molecular size markers are added to each side of the gel at 0.4 pg per well. v. Run the gel for one hour at 120 volts then stain in a 1% ethidium bromide solution for 20 min. Destain the gel for 45 min in deionized water and view the gel on a UV illuminated light box. Photograph gel to record results.

Duck plague PCR interpretation: A 500 bp amplification band in the lambda control sample indicates the PCR ran successfully. A 446 bp band in the duck plague known DNA control indicates the duck plague primers are working. A 446 bp band in the unknown test sample indicates duck plague viral DNA was present. No amplification products will be present in the duck plague or lambda DNA controls. If bands appear in these negative control products, cross contamination occurred during the and the test must be repeated. Alternative methods to detect DVE virus DNA have been published by Plummer et al. (14), and Pritchard et al. (15)

between 0 and 1.5 were detected in domestic and wild waterfowl that had not been exposed to DVE; an NI of 1.75 or greater was considered as evidence of prior exposure to DVE virus (3). Alternatively, sera may be screened using a constant-virus varyingserum method. In the author’s laboratory, a microtiter neutralization assay using primary MDEF or DEF cells is used. Serial two-fold dilutions of each serum sample (heat inactivated at 56 C) are prepared in 50 μΐ of serum-free EMEM in microtiter plates. Approximately 102 0 mean tissue culture infective dose (TCID50) of DVE virus in 50 μΐ of EMEM are added to each well and the mixtures are allowed to react at 37 C for 1 hr. A suspension of primary MDEF or DEF cells in EMEM supplemented with 2 mM

SEROLOGIC DETECTION IN THE HOST Serologic detection of infection is only of importance in noncommercial waterfowl and may be used to determine prior exposure of birds to DVE virus. The humoral response to natural infection with DVE virus is often low and antibodies may be short-lived (6); cell mediated immunity is assumed to also play a role in the infection (16). However, detection of neutralizing antibodies to DVE virus in serum is possible. VN assays, using a constant serum varying virus method, may be performed in chick or duck embryos by using embryoadapted virus, or in cell cultures. Neutralization indices (NIs) 108

Chapter 23

L-glutamine, 0.17% sodium bicarbonate, and 10% FCS is adjusted to contain 3 x 105 cells/ml. Cells are next added to the plates at 100 μΐ per well and the plates are then incubated for up to 96 hr at 37 C in a humidified 5% CO2 atmosphere. Following incubation, cultures are observed daily by light microscopy and finally fixed with 10% formol-buffered saline and stained with 1% crystal violet. The plates are read macroscopically. The titer for virus-neutralizing activity is expressed as the reciprocal of the highest dilution of serum at which there is no evidence of cytopathic effects and therefore complete virus neutralization has occurred. A titer of less than 3 log2 is usually considered negative. A VN titer of 8 or greater is considered significant and is evidence of exposure to DVE virus (6). VN antibody may also be detected using cell cultures by mixing sera at a single dilution (e.g., 1:10) with 100200 TCID50 virus and then testing inoculated cell cultures for non­ neutralized virus by immunofluorescence. Although this method is not quantitative, it can be useful for screening large numbers of sera. These latter methods, using constant virus and varying serum, require less sera than the NI methods.

Duck Vine

5. Deng, Μ Y., E. C. Burgess, and T. M Yuill. Detection of dock ρΜρκ virus by reverse passive hemagglutination test Avian Dis. 28 616-62X. 1984. 6. Docherty, D. E., and C. J. Franson. Duck Virus Enteritis. In: Vi ii i··^ Diagnostic Virology, A. E. Castro and W. P. Heuschele, eds. Mosby Yor Book, St Louis, Missouri, USA. pp. 25-28. 1992. 7. Erickson, G. A., S. J. Proctor, J. E. Pearson, and G. A Giwtrfw Diagnosis of duck virus enteritis (duck plague). 17th Annual Proceedings of the American Association of Veterinary Laboratory Diagnostic^·!. AAVLD, Madison, Wisconsin, USA Roanoake, Virginia, USA pp. 85-90L 1974. 8. Gough, R. E., and D. J. Alexander. Duck Virus Enteritis in Greai-Brii··, 1980 to 1989. Vet. Rec. 126:595-597. 1990. 9. Hansen, W. R, S. E. Brown, S. W. Nashold, and D. L. Knudso·. Identification of duck plague virus by polymerase chain reaction. Avian Dti 43:106-115. 1999. 10. Hansen, W. R., S. W. Nashold, D. Docherty, E., S. E. Brown, and D. L Knudson. Diagnosis of Duck Plague in Waterfowl by Polymerase Cham Reaction. Avian Dis. 44:266-274. 2000. 11. Islam, M R., J. Nessa, and K. M Halder. Detection of duck plague virus antigen in tissues by immunoperoxidase staining. Avian Path. 22:389393. 1993. 12. Kisary, S., and L. Zsak. Comparative studies on duck viral enteritis (DVE) virus strains in geese. Avian Path. 12:395-408. 1983. 13. Kocan, R. M Duck plague virus replication in Muscovy duck fibroblast cells. Avian Dis. 20:574-580. 1976. 14. Plummer, P. J., T. Alefantis, S. Kaplan, P. O'Connell, S. Shawky, and K. A. Schat. Detection of duck enteritis virus by polymerase chain reaction. Avian Dis. 42:554-564. 1998. 15. Pritchard, L. I., C. Morrissy, K. Van Phuc, P. W. Daniels, and H. A Westbury. Development of a polymerase chain reaction to detect Vietnamese isolates of duck virus enteritis. Vet. Microbiol. 68:149-156. 1999. 16. Richter, J. Η. M, and M. C. Horzinek. Duck Plague. In: Virus Infections of Birds, J. B. McFerran and M S. McNulty, eds. Elsevier Science Publishers B.V., Amsterdam, the Netherlands, pp. 77-90. 1993. 17. Sandhu, T. S., and S. A Shawky. Duck Virus Enteritis (Duck Plague). In: Diseases of Poultry, 11th ed. S. Y.M, H. J. Barnes, J. R. Glisson, A M Fadly, L. R. McDougald and D. E. Swayne, eds. Iowa State Press, pp. 354363. 2003. 18. Shawky, S., and K. A. Schat. Latency sites and reactivation of duck enteritis virus. Avian Dis. 46:308-313. 2002. 19. Wolf, K., C. N. Burke, and M C. Quimby. Duck viral enteritis: a comparison of replication by CCL-141 and primary cultures of duck embryo fibroblasts. Avian Dis. 20:447-454. 1976.

DIFFERENTIATION FROM CLOSELY RELATED AGENTS

DVE is usually pathologically distinct from other diseases of waterfowl; however, hemorrhages and necrosis do occur in other diseases of waterfowl such as duck viral hepatitis, pasteurellosis {Pasteurella multocida) and some toxic conditions. Occasionally, lesions caused by DVE virus could be confused with those produced by highly pathogenic Newcastle disease virus, avian influenza virus, and poxvirus (16, 17). REFERENCES

1. Brand, C. J. Duck Plague. In: Field Guide to wildlife diseases, Vol. 1: General field procedures and diseases of migratory birds, M Friend, ed. Resources publication 167. U.S. Department of the Interior, Fish and Wildlife Service, Washington, DC. pp. 117-127. 1987. 2. Burgess, E. C., and T. M Yuill. Increased cell culture incubation temperatures for duck plague virus isolation. Avian Dis. 25:222-224. 1981. 3. Dardiri, A H., and W. R. Hess. The incidence of neutralizing antibodies to duck plague virus in serums from domestic ducks and wild waterfowl in the United States of America. Proceedings, Annual Meeting of the United States Animal Health Association, pp. 225-237. 1967. 4. Dardiri, A H., and W. R. Hess. A plaque assay for duck plague virus. Can. J. Comp. Med. 32:505-510. 1968.

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24 HERPESVIRUSES OF FREE-LIVING AND PET BIRDS Erhard F. Kaleta SUMMARY. Free-living and pet birds are affected by a large number of different herpesviruses (HVs). The taxonomic status within the three subfamilies α-, β-, and y-Herpesvirinae is still undetermined for most of the many isolates. HV isolates can be distinguished by restriction enzyme analysis, cross neutralization tests, and in part by plaque morphology in chick-embryo fibroblast (CEF) and chick embryo kidney or chick embryo liver cell cultures. Replicating and latently present HVs can be detected in swabs and various tissues by polymerase chain reactions (PCRs). Most commonly affected by HVs are various breeds of domestic pigeons, waterfowl, many species of psittacines, owls, black and white storks, and various finches; eagles, falcons, cormorants, toucans, and quail are less frequently infected. Subclinical infections occur in all species. Various forms of apparent disease are frequently linked to the presence of debilitating factors such as overcrowding, food primarily deficiencies, prolonged antibacterial treatment, and poor hygiene. In the acute phase of the disease lesions are primarily hemorrhagic and inflammatory in nature and in the chronic phases lesions are predominantly necrotic mostly in the pharynx, esophagus, and intestines, but also in large parenchymal organs and brains. Agent Identification. Diagnosis of HV infections in live birds is preferably made by virus isolation from buffy coat cells and feather pulp material. Homogenates of parenchymal organs, bone marrow, or brain tissues from dead birds may be used as inocula for cell cultures of avian origin. Viral DNA can be detected by PCR. Virus identification is achieved by chloroform treatment of supernatant fluids from heavily infected cell cultures, cultivation of the isolate in question in the presence of iododeoxyuridine, and electron microscopy. Identification of serotypes is done by cross neutralization tests. Serologic Detection in the Host. Subclinical infection is commonly associated with seroconversion. The assay of sera in neutralization tests identifies infected, possibly also virus positive birds. In many cases several serologically unrelated viruses must be used to detect all serotypes of HVs known to occur in a particular species of bird. Repeated serologic monitoring is advisable for all bird collections, breeding stations, and quarantine stations. and 4 (31,64). Transmission experiments that prove the etiology and genesis of these papillomatous tumors have.not been reported; 2. the hemorrhagic type in infectious laryngotracheitis (ILT) infections in chickens and in pheasants (11,13,41), peafowl (3,5,13,42), and in guinea fowl (75); and 3. the necrotic type, which is found in many birds dying of HV infections. These birds include owls (6), falcons (51,74), eagles (16), rural and fancy pigeons (63,71) many psittacines (30,45,50,54), black storks (Ciconia nigra; 38) and white (Ciconia ciconia) storks (39), quail (40), and cranes (7), as well as passeriforms of the families Estrildidae (exotic finches), Ploceidae (weavers), and Carduelidae (finches, including the common canary). Respiratory distress, hepatitis, and disseminated focal necrosis located in liver, spleen, and bone marrow are most commonly seen (61,68,72,76).

INTRODUCTION Domestic, free-living, and pet birds are frequent hosts of a large variety of herpesviruses (HVs). Infections may go unnoticed or may result in fatal disease. HVs often interfere with captive breeding programs or with attempts to re-establish free-living bird populations. In addition, enzootics may occur in wild birds. CLINICAL DISEASE

More than any other viruses, HVs have a tendency for latency and persistent infections. Young age, concurrent infections, and noninfectious noxae may precipitate disease, whereas antibacterial, antifungal, and antiparasitic treatments may ameliorate the course of the disease. In any case, clinical signs appear primarily as nonspecific, general depression of common behavioral traits (46). Psittacines often have yellowish droppings. However, in a large number of cases, sudden increases in mortality rates may be the only signalment evident in many species of birds. Therefore, any increased death rate should prompt a detailed examination (18). In psittacines, HV-induced mortality is often preceded by introductions of latently infected birds, such as South American conures of the genera Pyrrhura and Cyanoliseus (22). These birds tend to be more often latently infected and are prolonged shedders than are other psittacines (21,23). Falcons, and in particular owls, often acquire fatal infections from preying on persistently infected domestic pigeons (2). Housing of large numbers of birds of various species in quarantine stations and pet bird shops provide ideal circumstances for the horizontal spread of different viruses, including HV, and also avian paramyxoviruses (10,25,34). Gross lesions in birds dying from HV-induced diseases (17,24,32) allow categorization of the disease into three main types: 1. the neoplastic type in chickens due to serotype 1 Marek’s disease viruses and in pigeons due to an HV antigenically unrelated to Marek's disease or classical HV infection of pigeons. Circumstantial evidence for a causal relationship between mucosal tumors in neotropical parrots and HVs has been reported (47,55). These mucosal tumors in pharynx and cloaca of neotropical parrots of the genera Ara and Amazona are histomorphologically similar to papillomas and are linked to psittacid HVs of genotypes 1, 2, 3,

SAMPLE COLLECTION Latent, non-productive infections are difficult to detect, and cyclophosphamide treatments before sampling may enhance recovery of the agent. During pairing time in seed-eating birds that feed their offspring with crop-milk and/or macerated seeds (e.g., pigeons and some psittacines, but also many passerine species), HV infections change from the latent, non-productive state to the persistent, productive state. Swabs of the oropharynx should be placed in complete tissue culture medium without serum. This medium is suitable for storage of samples at 4 C for about 1 wk. If necessary, infectivity in swabbed epithelial cells can be preserved in freezing medium; (80 ml basal medium, Eagle’s [BME] or M199, 10 ml fetal calf serum [FCS], and 10 ml dimethylsulfoxide [DMSO]; or in sucrosephosphate-glutamate albumin [SPGA] buffer) (8). In addition to swabs, tissue specimens should be taken from all organs with macroscopically visible lesions, (e.g. liver and spleen) in birds that die from suspected HV disease. Bone marrow specimens are valuable if carcasses have advanced autolysis. Specimens should be minced immediately and immersed in cell­ culture medium with twice the normal strength of antibiotic and antimycotic drugs. Such samples can be kept refrigerated for up to 1 wk without appreciable loss of infectivity. Freezing of samples should be avoided. If long-term storage is necessary, tissue homogenates should be suspended in freezing medium (see above).

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Stork HV induce a long-lasting viremia, with the bulk of HV infectivity associated with peripheral leukocytes (39). Heparinized whole blood is aseptically drawn from the wing vein of storks and further processed for virus isolation, as in the case of Marek's disease herpesvirus (see Chapter 22 on Marek’s disease).

Herpesviruses of Free-living and Pei Bmk

Characterization Culture filtrates that are free of bacteria, mycoplasmas, chlamydiae, and fungi and that induce any of the three types of cellular focal alterations described above should be examined further to confirm the presence of HVs. The type of lesions in affected birds, the tissues that yield isolates, and the types of cellular alterations are an indication, but not proof, of the presence of HVs. Virus identification involves physicochemical and biological properties, and, finally, immunologic identification. Molecular characterization, e.g. in situ hybridization (ISH), restriction fragment length polymorphism (RFLP) and several techniques of polymerase chain reaction (PCR) have gained increasing importance.

PREFERRED CULTURE MEDIA AND SUBSTRATES

Cell cultures are the preferred substrate and any common cell culture medium (e.g., BME, minimum essential medium [MEM] and modifications, or Ml99) that supports the growth and maintenance of cultured avian embryonic cells or young chick cells is suitable for the isolation of avian HVs (see Chapter 43 on cell culture methods). DNA of HVs that are in the non-productive stage (latency) can be detected by various PCRs. All known avian HVs multiply well in primary monolayer cultures of chicken kidney cells (CKCs) and also chick embryo liver cells (CELC). Marek's disease, ILT, and Lake Victoria Cormorant HV grow only in chicken kidney epithelium or CELC. It is laborious to adapt these three viruses to chick embiyo fibroblasts (CEFs). Other cells (e.g., duck embiyo fibroblasts or cell lines of mammalian origin) either are not definitely free from adventitious agents or yield erratic results. Therefore, such cell cultures cannot be recommended. Cells may be inoculated immediately after plating or after the formation of subconfluent monolayers. Incubation for more than 1 wk is not advisable. Instead, inoculated trypsinized cells should be subcultured at intervals of 4-6 days. Depending on the particular strain of HV, three types of cellular alterations can be observed in unstained cultures: 1) Small plaques consisting of roundish refractile cells on top of otherwise normal-appearing confluent CKCs or CEFs. 2) Large plaques with a lytic center and rounded cells adjacent to the periphery of the plaque. With extended incubation, more and more rounded cells detach and float in the medium. In the final stages a netlike structure of unchanged elongated cells remains. 3) Syncytial-type plaques composed initially of large irregular structures. Upon further incubation, these syncytia detach from the surface of the culture vessel and assume roughly spherical shapes and float in the medium and later disintegrate into particulate matter with irregular boundaries and different diameters. Some avian HV isolates may contain small and large plaque variants (35). An inoculum can be considered to be negative if the primary culture and two subsequent subcultures do not yield any of the described cytopathologic changes. Most HV isolates can be kept frozen without detrimental effects on viral titer. However, most of the stork isolates (39) and at least some of the strains recovered from psittacines and pigeons (33) are highly cell-associated. Preservation of such strains requires the addition of SPGA buffer (8,19) before freezing.

Physicochemical Properties Essential Lipid. All HVs are sensitive to lipid solvents. HVs enclosed in syncytia may escape the direct action of chloroform, leaving some intact particles unaffected. Centrifugation or filtration through 220-nm filters before chloroform treatment eliminates this problem. Filtration. An estimate of the average diameter of virions can be obtained by sequential filtration sets. Iododeoxyuridine. Virus replication is inhibited, but not entirely prevented, by halogenated desoxyribonucleosides. Morphology. Cell lysates of heavily inoculated cultures showing cytopathic effect (CPEs) are prepared either by one or two freeze­ thaw cycles or by sonication. After clarification by low-speed centrifugation, viral particles in the supernatant are pelleted by ultracentrifugation. The pellet is resuspended in saline, negatively stained, and examined in a transmission electron microscope. All complete enveloped HVs have the appearance of a fried egg, that is, with an irregular envelope and a nucleocapsid roughly in the center. Naked, nonenveloped particles or particles with disrupted envelopes may also be seen (9,10,24,45,59,62,69). Naked particles tend to form clusters consisting of 3-10 nucleocapsids. Because of relatively low infectivity titers (usually in the range of ΙΟ4—106 median tissue culture infective doses [TdD50]/ml), negative results are likely. In such situations, several steps of multiplication, purification, and concentration are needed. Because of variations in steps used for the processing of samples for electron microscopy and because of inherent differences in the examined isolates, diameters of the nucleocapsid vary between 100 nm and 150 nm and of the enveloped particles from 150 nm to more than 200 nm. Some HVs, such as several stork isolates, have an envelope that tightly surrounds the nucleocapsid. Others, such as owl, falcon, eagle, psittacine, crane, and quail HVs, carry a large, sometimes folded envelope (40). Spikes are not always clearly discernible on the outer surface of the envelope. Biological Properties Embryo Inoculation. Herpesviruses do multiply following inoculation of yolk sacs or chorioallantoic membranes (CAMs) in embryonating eggs from chickens or other birds. Inoculation by either route results in variable embryo mortality and, if the embryo survives the inoculation for more than 4 days, distinct pox-like lesions form on the CAM. The embryo itself may be stunted and have disseminated focal necrosis of the liver and spleen; occasionally focal lesions are present on the mucosa of the palatine bone. Allantoic fluid does not agglutinate chicken erythrocytes. Intranuclear Inclusions. Predominantly eosinophilic Cowdry Type A intranuclear inclusion bodies surrounded by a distinct halo can be detected in hematoxylin-and-eosin-stained sections of liver, spleen, and bone marrow cells of naturally infected birds (9,10,24,45,59,62,69); in cultured cells inoculated with avian HVs (9,12,15,34,37,42,51,62); and liver and spleen cells of infected embryos (6,18,33,48).

AGENT IDENTIFICATION Taxonomy Herpesvirus species of free-living and pet birds are currently not assigned to any of the three subfamilies of the α-, β-, and γHerpesvirinae (52,60). Moreover, names for the diseases in various bird species or higher taxons have not yet been coined. In this paper, names for HV isolates are selected according to suggestions made by Roizman (60). Table 24.1 presents an updated list of 27 known and sufficiently characterized HV grouped in chronologic order according to the first description of a defined disease or of the causative virus. Additional data refer to the main host species (or group), the provisional species designation (60), the serologic relatedness of the isolates to each other (37,40), and the predominant type of lesion in naturally infected birds (32). Ill

Erhard F. Kai eta

Animal Inoculation. Numerous in vivo studies with HVs derived from domestic birds have yielded valuable information on the development of signs, lateral spread and lesions and contributed to the knowledge of pathogenesis, prevention and immunoprophylaxis. Pathogenicity studies with avian HVs and naturally free-living and pet birds are generally of limited value. Maintenance of such birds in biocontainment isolation cabinets is difficult because of the birds’ sometimes extreme demands on the environment and food. The general health of such birds and their immune status to HVs are often unknown; their immunocompetence might be impaired. Legal rulings on animal protection and ethical reasons prevent the handling of endangered species. Data on the host range of HVs can be obtained more effectively by swabbing the pharynx and cloaca or by assaying purified peripheral leukocytes for viremia. Indirect evidence for the presence of natural infections by various HVs can be gathered by testing serum samples for neutralizing antibodies (15,28). The domestic chicken is not susceptible to any avian HVs other than Marek's disease HVs and ILT virus. Genomic properties. Parts of the genome of some psittacine and passerine HVs were studied (27,66,67) using consensus primer PCRs that amplify either a region in the DNA-directed DNA polymerase gene (70,76) or parts of the UL26 open reading frame (66,67). The UL6 and UL7 of duck enteritis HV were amplified by Plummer et al. (56). The results of these studies confirmed in most instances the genetic heterogeneity of psittacine HVs, the results of previous cross neutralization tests and earlier data on the restriction fragment length polymorphisms (26,27,34,37,40). Although assignment of the examined HVs to any of the three established subfamilies of the family Herpesviridae is not yet achieved, the genomic data contribute to the phylogenetic evolution of avian, mammalian and reptilian HVs (12,14).

SEROLOGIC DETECTION IN THE HOST Neutralizing antibody in sera of birds can be detected by the plaque-reduction method in primary CEF or CKC cultures. Thirty to 100 plaque-forming units of virus are reacted with twofold dilutions of antisera and incubated for at least 1 hr at room temperature. Two plates are inoculated as negative controls. All cultures are stained with neutral red (2 ml of a 1:40,000 dilution in phosphate-buffered saline per 60-mm petri dish culture) after 5-6 days of incubation and examined macroscopically for plaques. Plaque numbers are expressed as a percentage of negative controls. The serum dilution is then plotted against the probit values, and the titer of the antisera is estimated graphically and enumerated as log2 values. A less laborious neutralization test can be performed under less stringent conditions in 96-well microtiter plates. Twofold serum dilutions ranging from 1:2 to 1:256 are incubated with 100 TCID50 of specific HV for at least 1 hr. Freshly prepared CEF at a concentration of 106 cells/ml are added. After 5-6 days of incubation, plates are stained with crystal violet solution, and serum titers are macroscopically enumerated. The titer of a serum is the highest dilution that produces no CPEs. Plaques also can be enumerated by direct or indirect immunofluorescence (57,58). Other serologic tests, such as agar gel precipitation, complement fixation, or enzyme-linked immunosorbent assay are either insufficiently sensitive or not yet developed and validated (3). DIFFERENTIATION FROM CLOSELY RELATED AGENTS

Using the neutralization test, some avian herpesviruses are shown to be serologically closely related (Table 24.1). These are Marek's disease and turkey herpesviruses; HVs of pigeons, owls, falcons, and eagles; and HVs from cranes and bobwhite quail. No serologic relatedness is detectable among any of the other viruses listed in Table 24.1. It should be noted that at least five serologically different herpesviruses exist within birds of the order Psittaciformes (26). The psittacine HV of the serotype 1 is most frequently isolated. Also, two serologically distinguishable HVs can be recovered from pigeons. The serologic status of HVs recently isolated from passeriforms in this laboratory (exotic finches, weaver bird, and common canaiy) indicates a relationship to psittacine viruses (26,27,68). Any enhanced morbidity and above-normal mortality in free-living and pet birds requires a comprehensive postmortem and laboratory examination. Certain bacteria (e.g., Salmonella spp., Klebsiella spp., Escherichia coli, Mycobacterium spp., Campylobacter spp., Pseudomonas spp., Bacillus spp., Erysipelothrix rhusiopathiae, and Pasteurella spp.) or bacterial exotoxins (e.g., botulism) may cause focal necrosis in livers. Hemorrhagic lesions in the respiratory tract might be caused by certain bacterial infections (e.g., Pasteurella spp.) or by the red worm Syngamus trachea. Highly pathogenic avian influenza and Newcastle disease viruses grow readily in cell cultures and embryonating eggs. Members of both groups of viruses easily can be detected by hemagglutination in cell culture and allantoic fluids of inoculated eggs. Diphtheritic lesions in the intestines might be caused by Trichomonas spp. if the organisms are found in the pharynx or esophagus. Jejunal diphtheritic lesions and hemorrhages can be induced by Clostridium spp. Chlamydophila (formerly Chlamydia) psittaci, as a possible cause of respiratory distress, is widespread in various birds (42,46). Its detection is possible with cell culturing and other techniques. Laboratory workers must take special precautions to avoid becoming infected. Several avian viruses that also induce CPEs and/or intranuclear inclusions need to be excluded (49,70). These are polyomaviruses

Immunologic Identification Antisera against specific HVs can be produced in rabbits and used with the plaque-reduction method in primary CEF or CKC cultures (see serologic detection in the host below) to categorize unknown HV isolates into serogroups. Avian HVs are poorly immunogenic in laboratory rodents. The following procedure has been adopted to raise antisera in rabbits (26,27). Viruses are harvested, and cells are disrupted by ultrasonic vibration. Cellular debris is removed by centrifugation at 4000 x g for 30 min. The supernatant fluid is mixed with 10% (w/v) polyethylene glycol MW 6000, stirred at 4 C, and kept overnight (48). After centrifugation at 4000 x g for 10 min, the pellet is resuspended in Dulbecco’s phosphate buffer and layered on top of a 12-52% linear sucrose gradient as described by Lee et al. (48) and centrifuged at 25,000 x g for 1 hr in a Beckman SW 27 rotor (Beckman Instruments, Inc, Fullerton, Calif.). The gradient fluid is collected in 2-ml fractions. The same fractions are titrated in cell cultures for infectious virus. Fractions with the highest viral content are mixed in equal parts of complete Freund's adjuvant and are inoculated intradermally on multiple sites on the back of a rabbit. The first antiserum is collected 4 wk later. Booster injections are given at 2-wk intervals. Rabbits are bled 2 wk after the third booster injection is given. Serum can be collected and stored until used for serotyping unknown HVs. Using the plaque-reduction method in primary CEF or CKC cultures, 11 different serotypes (33,37) can be distinguished. In Table 24.1, available data on the serologic relatedness of herpesviruses is indicated in the fourth column. Available, yet preliminary, data on DNA restriction endonuclease analysis (2,27) seem to confirm the results of conventional neutralization tests. The use of monoclonal antibodies for identification and differentiation of avian HVs is hampered by the large number of different viruses from various bird species. So far only one report exists on studies of a psittacine HV (1). 112

Chapter 24

Herpesviruses of Free-living and Pet Birds

(65). Biological substances such as various preparations of propolis and several caffeoylics inhibit the multiplication in vitro and in vivo of HVs from pigeons and psittacines (35,44). All these drugs need to be given very early and repeatedly after infection. Local reactions at the site of injection may occur. Several attempts have been made to protect psittacines by adjuvanted vaccines containing partially purified, formolinactivated viruses (20,21,29,36,77). The widespread prophylactic use of such vaccines is limited by several factors. These include the broad antigenic diversity of psittacine HVs (26,66,67), which precludes the use of one seed virus for vaccines in all cases and undesirable side effects such as local inflammatory or granulomatous to necrotic reactions at the site of inoculation. In addition, strict but variable governmental regulations for such ’’niche" products in various countries makes it cumbersome for commercial enterprises to develop and produce these vaccines. Available information on vaccination-challenge experiments or seroconversion are promising and favor additional studies on innocuity, immunogenicity, and duration of protection.

(especially psittacines), reoviruses, rotaviruses, and adenoviruses (in many species, particularity in pigeons); paramyxoviruses (in almost all avian species); orthomyxoviruses (frequent in waterfowl and gallinaceous birds); togaviruses and coronaviruses (in turkeys); and also poxiruses (4). Viruses that might be present but do not necessarily cause CPEs include picomaviruses (in waterfowl and gallinaceous birds), parvoviruses (in geese and Muscovy ducks), and some retroviruses (in ducks and turkeys), in psittacines); Control and vaccination. All HVs are horizontally transmitted via saliva or droppings. The detection of shedders by virologic examination of swabs taken from the oropharynx and cloaca (see above) and separation of shedders from noninfected susceptible birds greatly helps to reduce fatal infections (23). Cell-free viruses in the environment are sensitive to inactivation by ultraviolet light (sunshine) and also heat and chemical disinfectants (73). The application of inhibitors of viral DNA synthesis such as acyclovir and gancyclovir resulted in reduced rates of morbidity and mortality following experimental exposure of Quaker parakeets (53). Some strains of pigeon HVs also are sensitive to acyclovir Table 24.1. Updated list of avian herpesviruses (Hvs).

No. 1

2 3

4 5 6 7 8 9 10 11

12 13 14

15

Name of disease/infection Marek s disease Turkey HV infection Duck viral enteritis Infectious laryngotracheitis Pacheco's disease Pacheco's disease Pacheco's disease Pacheco's disease Pacheco's disease Smadel 's disease Pharyngo esophagitis Hepatosplenitis infectiosa strigum Inclusion body hepatitis Inclusion body hepatitis

16 17

None Inclusion body hepatitis None

18

None

19 20 21 22 23 24 25 26 27

None None None None None None None None None

Susceptible bird (species)/ HV isolated from Chicken

Provisional HV species designationA GallidHV2

Serologically related to no. 2

Type of lesion Tumorous

Turkey, chicken Ducks, geese, swans Chicken, pheasants, others Psittaciformes Psittaciformes Psittaciformes Psittaciformes Psittaciformes Pigeons

Meleagrid HV 1 AnatidHVl

1 None

None Hemorrhagic

GallidHVl Psittacid HV 1 Psittacid HV 2 Psittacid HV 3 Psittacid HV 4 Psittacid HV 5 ColumbidHVl

None 20-23, 25 None None None 19 12,13,14

Hemorrhagic Necrotic Hemorrhagic Necrotic Necrotic Necrotic Necrotic

26,61 26,44 26, 33 26 26 11,52

Pigeons

Columbid HV 2

None

Necrotic

33

Various owls

StrigidHVl

10,13,14

Necrotic

6

Falcon

FalconidHVl

10,12, 14

Necrotic

50, 73

Bald eagle nestling Lake Victoria cormorant

Acciprid HV 1 Phalacrocoracoid HV 1

10,12, 13

Necrotic

16

None

None®

19

Cranes Bobwhite quail Black and white storks Black-footed penguin Exotic finch Weaver bird Canary Exotic finch Exotic finch Tragopan Superb starling Toucan

GruidHVl PercididHVl

17 16

Necrotic Necrotic

7, 18 40

Ciconiid HV 1

None

Necrotic

34,39

Sphenicid HV 1 EstrildidHV 1 PloceidHVl Serinid HV 1 EstrildidHV 2 EstrildidHV 3 Tragopanid HV 1 Lamprotomid HV 1 Andigenid HV 1

4? 9 5 5 5 5 None 5

Hemorrhagic Hepatitis Hepatitis None® None® None® None® Hepatosplenitis Necrotic

43 60,71,76 60,71 27 27 27 27 27 9

A Modified from Roizman (49). B No lesions observed so far

113

9

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Erhard F. Kaleta

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65. Thiry, E., H. Vindevogel, P. Leroy, P.-P. Pastoret, A. Schwers, BL Brochier, Y. Anciaux, and P. Hoyois. In vivo and in vitro effect of acydovir on pseudorabies virus, infectious bovine rhinotracheitis virus and pigeon herpesvirus. Ann. Rech. Vet. 14:239-245. 1983. 66. Tomaszewski, E. K., G. van Wilson, W. L. Wigle, and D. N. Phaten Detection of hetero-geneity of herpesviruses causing Pacheco’s disease b parrots. J. Clin. Microbiol. 39:533-538. 2001. 67. Tomaszewski, E. K., E. F. Kaleta, and D. N. Phalen. Molecular phylogeny of the psittacid herpesviruses causing Pacheco’s disease correlation of genotype with phenotypic expression. J. Virol. 77:1126011267. 2003. 68. Tomaszewski, E. K., M Gravendyck, E. F. Kaleta, and D. N. Phalen. Genetic characte-rization of a herpesvirus isolate from a superb starling (Lamprotornis superbus) as a psittacid herpesvirus genotype 1. Avian Dis. 48:212-214. 2004. 69. Tsai, S. S., J. H. Park, K. Hirai, and C. Itakura. Herpesvirus infections in psittacine birds in Japan. Avian Pathol. 22:141-156. 1993. 70. Van Deventer, D. R., P. Warrener, L. Bennett, E. R. Schultz, S. Coulter, R. L. Garber, and T. M Rose. Detection and analysis of diverse herpesviral species by consensus primer PCR. J. Clin. Microbiol. 34:1666-1671. 1996. 71. Vindevogel, H., L. Dagenais, B. Lansival, and P. P. Pastoret. Incidence of rotavirus, adenovirus and herpesvirus infection in pigeons. Vet. Rec. 109:285-286. 1981. 72. von Rotz, A., A. Ruebel, F. Mettler, and R. Hoop. Letal verlaufende Herpesvirusinfektion bei Goldsamadinen (Chloebia Gouldinae [Gould]). Schweiz. Arch. Tierheilkd. 126:651-658. 1984. 73. Wagner, U. Vergleichende Untersuchungen zur Empfindlichkeit verschiedener aviarer Herpesvirusisolate gegeniiber chemischen Desinfektionsmitteln. Veterinarxy Medical Thesis. Giessen, Germany. 1993. 74. Ward, F. P., D. G. Fairchild, and J. V. Vuicich. Inclusion body hepatitis in prairie falcon. J. Wildl. Dis. 7:120-124. 1971. 75. Watanabe, T., and H. Ohmi Susceptibility of guinea fowl to the virus of infectious laryngotracheitis and egg-drop syndrome—1976. J. Agric. Sci. Tokyo Nogyo Daigaku 28:193-200. 1983. 76. Wellehan, J. F. X., M Gagea, D. A. Smith, W. M Taylor, Y. Berhane, and D. Bienzle. Characterization of a herpesvirus associated with tracheitis in Gouldian finches (Erythrura [Chloebia} gouldiae). J. Clin. Microbiol. 41:4054-4057. 2003. Π. York, S. M, and C. J. York Pacheco virus vaccine studies—A preliminary report. In: Proceedings of the 32rd Western Poultry Disease Conference, Davis, Calif. February 8-10. pp. 101-102. 1983.

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25 POX Deoki N. Tripathy and Willie M. Reed

SUMMARY. Avian pox is a common viral disease of domestic birds (chickens, turkeys, pigeons, canaries) and wild birds. Approximately 232 species in 23 orders of birds have been reported to acquire natural poxvirus infection. Avian pox is a slow-spreading disease characterized by the development of proliferative skin lesions (cutaneous form) and/or upper digestive and respiratory tract lesions (diphtheritic form). The causative agent is a double-stranded DNA virus of the genus Avipoxvirus of the family Poxviridae. Agent Identification. Pox is routinely diagnosed by histologic examination of proliferative skin, oral, or tracheal lesions for cytoplasmic inclusion bodies in tissue sections, or viral particles exhibiting typical poxvirus morphology using electron microscopy. The virus is isolated by inoculation of chorioallantoic membranes (CAMs) of 9 to 12-day embryonating chicken eggs. Pocks develop on the CAM in 5-7 days. The etiology is confirmed by histopathologic or ultrastructural examination of the CAM lesions for cytoplasmic inclusions or viral particles with typical poxvirus morphology, respectively. Immunoperoxidase or indirect fluorescent antibody and agar-gel immunodiffusion (AGID) tests can also be used to detect pox viral antigen in tissue samples. Antigenic differences among isolates can be determined by immunoblotting, host pathogenicity, and virus neutralization. Isolated viruses can be evaluated for pathogenicity for susceptible hosts or for susceptibility in avian cell culture. Molecular characterization is done by restriction fragment length polymorphism (RFLP) analysis of viral genome, polymerase chain reaction (PCR) amplification of specific genomic fragments and nucleotide sequence determination of the genome or selected fragments. Serologic Detection in the Host. Serologic detection of infection may be important in experimental studies and for measuring the immune responses following vaccination. Antibody responses can be measured by AGED, passive hemagglutination, immunoperoxidase, and enzymelinked immunosorbent assay (ELISA). Monoclonal antibodies against fowlpox have been used in experimental studies to differentiate strains of fowlpox virus. Modified live virus vaccines (fowl pox, pigeon pox, canary pox, turkey pox, and quail pox viruses) of chick-embryo or cell-culture origin are available commercially. becomes yellowish brown to dark brown. Removal of such lesions before they are not completely dry leaves a hemorrhagic, moist surface. When the scab is dry it drops off, leaving a scar. In the diphtheritic form of pox, lesions occur on the mucous membrane of the mouth, nares, pharynx, larynx, esophagus, or trachea. Mouth lesions can interfere with feeding. Tracheal lesions can cause difficulty in breathing and may simulate signs of infectious laryngotracheitis in chickens. In layers, the disease causes a drop in egg production, and in young chicks it reduces growth. Death occurs in cases with the generalized infection or diphtheritic form of the disease. The acute systemic disease caused by canary poxvirus results in high mortality in susceptible canaries. Although all avian species appear susceptible to pox (29), only fowl poxvirus has been studied extensively.

INTRODUCTION Approximately 232 species in 23 orders of birds have been reported to acquire a natural poxvirus infection (2). Although the number of distinct causative agents is unknown, all of them belong to the genus Avipoxvirus of the family Poxviridae. This genus includes fowl, turkey, pigeon, canary, and quail viruses, as well as other avianpox viruses. Fowlpox virus is a common pathogen of chickens and turkeys. The infection is generally manifested as either cutaneous or diphtheritic forms, although both types of the disease may occur in the same bird. In addition, an acute systemic form of pox is seen in canaries and results in high mortality. All forms of the disease (acute systemic, cutaneous, and diphtheritic) occurring either singly or in combination have been observed in canaries (4). In the cutaneous form, proliferative lesions are primarily confined to unfeathered areas of skin (e.g., legs, head, and eyelids), whereas in the much more lethal diphtheritic form, the lesions are found in the mouth, esophagus, and trachea. The acute systemic form of pox in canaries is characterized by fibrinous inflammation of serous membranes, pulmonary edema, and fibrinous pneumonitis. Because of the economic importance of this disease in commercial poultry, fowl pox virus and pigeon pox virus vaccines of chorioallantoic membrane (CAM) or cell-culture origin have been used for more than 60 yr. In recent years, turkey, canary and quail virus vaccines have also become available commercially. In spite of regular vaccination, outbreaks of fowlpox have occurred in all regions of the US in previously vaccinated chicken flocks.

SAMPLE COLLECTION Poxviruses are readily isolated from the nodular lesions of infected birds. Tissues with lesions, preferably recently developed lesions, can be removed with sterile scissors and forceps by cutting deep into the epithelial tissue. The material is ground with either sterile fine sand or 60-mesh aluminum oxide (Norton Alundum; Fisher Scientific Co., Pittsburgh, PA) with a sterile mortar and pestle or in a glass grinder. Hanks’ balanced salt solution, saline solution, or broth is added to make a 10% suspension. The suspension is centrifuged for 10 min at about 700 x g to remove large tissue particles. Antibiotics (penicillin and streptomycin) are added to the supernatant to give respective final concentrations of 1000 IU/ml and 1 mg/ml, and the suspension is held at room temperature for 30 min to 1 hr before inoculation. A piece of tissue should also be collected (in 10% neutral buffered formalin) for histopathologic examination to reveal cytoplasmic inclusion bodies or for transmission electron microscopy (TEM) to demonstrate virus particles by negative staining or in ultrathin sections. Serum samples may be collected to determine previous exposure by serologic tests.

CLINICAL DISEASE

The cutaneous form of pox is characterized by development of nodular lesions on various parts of unfeathered skin. A mild form of the disease may remain unnoticed, with only small focal lesions, usually on the comb and wattles. In the severe form of the disease, generalized lesions may occur on any part of the body, such as the comb, wattle, comer of the mouth, around the eyelids, the angle of the beak, the ventral surface of the wings, and the vent. Skin lesions may be small and discrete or may involve large areas through the coalescence of adjoining lesions. The surface of lesions is moist for a short time but dries soon, with a rough, irregular surface that

PREFERRED CULTURE MEDIA AND SUBSTRATES

Poxviruses can be isolated by inoculating the suspect material in

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embryonating eggs, susceptible birds, or cell cultures (e.g., chick embryo kidney cells, chick embryo dermal cells, or quail cells).

Pox

fluorescent antibody and immunoperoxidase methods (27). Viral antigen can be detected in lesions by indirect fluorescent antibody or immunoperoxidase tests using specific antiserum. A quick method for diagnosis of avian pox uses simultaneous fixation and dehydration and demonstrates cytoplasmic inclusion in less than 3 hr (17). Elementary bodies can be demonstrated in impression or squash-preparation smears stained by the Gimenez method (25). Negative-contrast TEM can be used for direct demonstration of viral particles in clinical material, pocks in CAM, or cultured cells (11).

Embryo inoculation The preferred and most convenient hosts are 9 to 12-day-old embryonating chicken embryos from a specific-pathogen-free flock. About 0.1 ml of the virus suspension is inoculated on the CAM. Inoculated embryos are incubated at 37 C for 5-7 days and then examined for pocks on the CAM. Pox lesions appear as either focal white opaque pocks or a generalized thickening of the CAM. Specific diagnosis requires either histologic demonstration of cytoplasmic inclusion bodies, or viral particles by TEM, or detection of specific poxvirus antigen in the CAM lesions. Occasionally, some strains from wild and pet birds fail to grow on the CAM and may require adaptation.

Antigen Detection in Tissues Poxviral antigens can be detected in cytoplasmic inclusions of infected cells (cutaneous or cultured) by direct or indirect immunofluorescent or immunoperoxidase microscopy using specific primary antiserum. Appropriate samples would include fowlpox-infected CAM, skin, or cell culture (27). The immunoperoxidase technique has the advantage of using a light microscope and sections can be stored for more than 2 mo without loss of staining.

Chicken inoculation The suspension made from a lesion is either applied to the comb, which is then scarified, or applied with a brush to denuded feather follicles (pluck 5 or 6 feathers). Similarly, wing web is another suitable site that can be used for inoculation. Susceptible chickens develop lesions at the site of inoculation in 5-10 days. Later, generalized lesions may appear during the disease, especially with virulent field strains. The disease is more severe in susceptible young birds than in older birds.

Molecular Identification Restriction fragment length polymorphism has been used for comparing closely related DNA genomes (7, 16, 24). In this regard, genomic profiles of quail, canary, mynah and poxviruses from Hawaiian endangered forest birds are different from those of fowl poxvirus (9, 24,28). Cloned genomic fragments of fowl poxvirus can be used effectively as nucleic acid probes for diagnosis (5). In this procedure, viral DNA isolated from legions is hybridized either with 32P-labeled or nonradioactive-labeled genomic probes. This method is especially useful for differentiation of the diphtheritic form of fowlpox from infectious laryngotracheitis when tracheal lesions are present (5).genomic DNA sequences of various sizes can be amplified by polymerase chain reaction (PCR) using specific primers (8, 10, 18, 24). This technique is very sensitive, especially when an extremely small amount of virus is present in the sample. Complete nucleotide sequence of a vaccine like fowlpox virus (1) and canarypox virus (30) has been determined. Molecular analysis reveals that while most strains of fowlpox virus from field isolates have integrated reticuloendotheliosis virus (REV) in their genome, the vaccine strains of fowlpox virus contain remnants of long terminal repeats (LTR) of REV (6, 18, 19, 23).

Cell culture Primary chick embryo, chick embryo kidney, chick embryo dermis, duck embryo, and secondary quail (QT-35) or LMH cells (16) support the growth of fowl poxvirus, producing a cytopathic effect (CPE), usually in 4-6 days, although CPE may occur earlier with high-titer inocula. The cells often become round and refractile, followed by degeneration. The virus produces plaques in agaroverlaid monolayers of chick embryo cells or quail (QT-35) cells. Plaques are produced in 3-5 days in QT-35 cells. Adaptation of strains to cell culture is necessary for plaque formation, because not all strains form plaques. The method has been considered unsuitable for titration of fowl poxvirus, because it is less sensitive than pock formation on the CAM (12).

AGENT IDENTIFICATION Physicochemical, Morphologic, and Biological Properties Fowl poxvirus, the type species of the genus Avipoxvirus, is a brick-shaped to rectangular virus. Other species of the genus are morphologically similar. Complete nucleotide sequence of the genomes of fowlpox virus and canary pox virus has been determined. The genome of fowl poxvirus is composed of a single double-stranded DNA molecule of approximately 288 kilobases (1). The genome of canary poxvirus is 365 kbp (30). Fowl poxvirus is resistant to ether. It is inactivated by 1% caustic potash when separated from its matrix. It withstands 1% phenol and 0.1% formalin for 9 days. Inactivation occurs by heating at 50 C for 30 min or 60 C for 8 min. Trypsin has no effect on the DNA or whole virus. Fowl poxvirus can survive in dried scabs for months or even years. In this regard, the presence of photolyase gene in the genome of fowlpox virus appears to play some role in virus persistence (21, 22). The virus multiplies in the cytoplasm of infected cells and forms inclusion bodies (Bollinger bodies) that contain the elementary bodies (Borrel bodies). It causes proliferation of the epithelium with ballooning degeneration of the cells and ultimately epithelial cell death. The cytoplasmic type-A inclusions vary in size and shape and can be demonstrated in histologic sections of skin, or in diphtheritic or CAM lesions. The inclusions can be stained by hematoxylin and eosin, acridine orange, and Giemsa stains and the Feulgen reactions. The specificity of the viral inclusions can be determined also by

SEROLOGICAL DETECTION IN THE HOST Agar Gel Immunodiffusion Test Precipitating antibodies can be detected by reacting test sera against viral antigens (25). The antigen can be derived by sonication and homogenization of infected skin or CAM lesions, as well as by treatment of infected cell cultures as described for ELISA. The lysed cell suspension is centrifuged and the supernatant is used as antigen. Gel-diffusion medium is prepared with 1% agar, 8% sodium chloride, and 0.01% thimersol. The viral antigen is placed in the central well and the test sera are placed in the peripheral wells. It is important to include a positive and negative control serum. The plates are incubated at room temperature. Precipitation lines develop in 24-48 hr after incubation of the antigen with antibody to homologous or closely related strains. The test is less sensitive than the ELISA (3) or die passive hemagglutination test (26, 29).

Passive Hemagglutination Antibodies against fowl poxvirus can be measured by a passive hemagglutination test using tanned horse or sheep erythrocytes (26) coated with fowl poxvirus antigen. Passive hemagglutinating antibodies are detectable in some sera of infected birds as early as 117

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1 wk after inoculation and may persist for 15 wk; that is, longer than precipitating antibodies. Cross-reactions occur among avian poxviruses.

REFERENCES

1. Afonso, C. L., E. R Tulman, Z. Lu, L. Zsak, G. F. Kutish, and D. L. Rock. The genome of fowlpox virus. J. Virol. 74:3815-3831, 2000. 2. Bolte, A. L., J. Meurer, and E. F. Kaleta. Avian host spectrum of avipoxviruses. Avian Pathol. 28:415-432, 1999. 3. Buscaglia, C., R. A. Bankowski, and L. Miers. Cell-culture virus­ neutralization test and enzyme-linked immunosorbent assay for evaluation of immunity in chickens against fowlpox. Avian Dis. 29:672-680, 1985. 4. Donnelly, T. M., and L. A Crane. An epomitic of avian pox in a research aviary. Avian Dis. 28:517-525,1984. 5. Fatunmbi, Ο. O., W. M Reed, D. L. Schwartz, andD. N. Tripathy. Dual infection of chickens with pox and infectious laryngotracheitis (ILT) confirmed with specific pox and ELT DNA DOT-BLOT hybridization assays. Avian Dis. 39:925-930,1995. 6. Garcia, Μ, N. Narang, W. M Reed, and A. M Fadly. Molecular characterization of reticuloendotheliosis virus insertions in the genome of field and vaccine strains of fowlpox virus. Avian Dis. 47:343-354,2003. 7. Ghildyal, N., W. M Schnitzlein, and D. N. Tripathy. Genetic and antigenic differences between fowlpox and quailpox viruses. Arch. Virol. 106:85-92,1989. 8. Kim, T. J., and D. N. Tripathy. Reticuloendotheliosis virus integration in the fowlpox virus genome: not a recent event. Avian Dis. 45:663-669,2001. 9. Kim, Tae-Joong, W. M Schnitzlein, D. McAloose, A. P. Pessier, and D. N. Tripathy. Characterization of an avian pox virus isolated from an Andean Condor Vultur gryphus. Veterinary Microbiology 96:237-246, 2003. 10. Lushow, D., T. Hoffman, and Η. M Hafez. Differentiation of avianpox virus strains on the basis of nucleotide sequences of the 4b gene fragment. Avian Dis. 48:453-463, 2004. 11. McFerran, J. B., J. K. Clarke, and W. L. Curran. The application of negative contrast electron microscopy to routine veterinary virus diagnosis. Res. Vet. Sci. 12:253-257,1971. 12. Morita, C. Studies on fowlpox viruses. Π» Plaque-neutralization test. Avian Dis. 7:93-98,1973. 13. Moyer, R. W., Β. M Arif, D. N. Black, D. B. Boyle, R. M Buller, K. R. Dumbell, J. J. Esposito, G. McFadden, B. Moss, A. A. Mercer, S. Ropp, D. N. Tripathy, and C. Upton. Family Poxviridae. In: Virus Taxonomy: Classification and Nomenclature of Viruses. Seventh Report of the International Committeee on Taxonomy of Viruses. Academic Press, San Diego, pp. 137-157,2000. 14. Reed, W. M, and Ο. O. Fatunmbi. Pathogenicity and immunological relationship of quail and mynah poxviruses to fowl and pigeon poxviruses. Avian Pathol. 22:395-400,1993. 15. Reed, W. M, and D. L. Schrader. Immunogenicity and pathogenicity of mynah pox virus in chickens and bobwhite quail. Poult. Sci. 68:631-638, 1989. 16. Schnitzlein, W. Μ, N. Ghildyal, and D. N. Tripathy. Genomic and antigenic characterization of avipoxviruses. Virus Res. 10:65-76,1988. 17. Sevoian, M A quick method for the diagnosis of avian pox and infectious laryngotracheitis. Avian Dis. 4:474-477,1960. 18. Singh, P., T. J. Kim, and D. N. Tripathy. Re-emerging fowlpox: evaluation of isolates from vaccinated flocks. Avian Pathol. 29:449-455, 2000. 19. Singh, P., W. M Schnitzlein, and D. N. Tripathy. Reticuloendotheliosis virus sequences within the genomes of field strains of fowlpox virus display variability. J. Virol. 77:585-586, 2003. 20. Singh, P., T.-J. Kim, and D. N. Tripathy. Identification and characterization of fowlpox virus strains using monoclonal antibodies. J. Vet. Diag. Invest. 15:50-54, 2003. 21. Srinivasan, V., W. M Schnitzlein, and D. N. Tripathy. Fowlpox virus encodes for a novel DNA repair enzyme that restores infectivity of UV-light damaged virus. J. Virol. 75:1681-1688,2001. 22. Srinivasan, V., and D. N. Tripathy. The DNA repair enzyme, CPDphotolyase restores the infectivity of UV-damaged fowlpox virus isolated from infected scabs of chickens. Veterinary Microbiology 108:215-223, 2005. 23. Tadese, T., and W. M Reed. Detection of specific reticuloendotheliosis virus sequence and protein from REV-integrated fowlpox virus strains. J. Virol. Meth. 110(l):99-104, June 9, 2003. 24. Tadese, T., and W. M Reed. Use of restriction fragment length polymorphism, immunoblotting and polymerase chain reaction in the differentiation of avianpox viruses. J. Vet. Diag. Invest. 15:141-150,2003. 25. Tadese, T., E. A. Potter, and W. M Reed. Development of a mixed agar gel enzyme assay (AGEA) for the detection of antibodies to poxvirus in chicken and turkey sera. J. Vet. Med. Sci. 65:255-258,2003.

Virus Neutralization Virus-neutralizing antibodies usually appear 1-2 wk after natural infection or experimental inoculation. The antibody response can be measured either by pock reduction in CAMs or plaque reduction in cell cultures (12). ELISA ELISA is a sensitive method to measure humoral antibody responses (3), and antibodies can be detected by 7 days after infection. Fowlpox virus antigens are prepared from either infected QT-35 cell monolayers or CAM lesions. Infected cells are pelleted (700 x g for 10 min at 4 C), washed with isotonic buffer (10 mM Tris, pH 8.0, 150 mM NaCl, 5 mM ethylenediamine tetraacetic acid [EDTA]) followed by lysis in hypotonic buffer (10 mM Tris, pH 8.0, 10 mM KC1, 5 mM EDTA) containing 0.1% Triton X-100 and 0.025% beta-mercaptoethanol. Nuclei and cellular debris are removed by low-speed centrifugation (500 x g for 5 min at 4 C) and the resulting supernatant is used as a source of fowlpox virus antigens for ELISA or immunoblotting. To isolate viral antigen from CAM lesions, initial grinding of the lesions with subsequent detergent treatment as described earlier would be required. Virus propagated in chicken embryo fibroblasts and chicken embryo dermis cells has also been used for antigen. Strain Variability Fowlpox virus is the type species of the Avipoxvirus genus and has been studied more extensively than other members of this genus (i.e., pigeon, canary, turkey, sparrow, junco, quail, mynah and other avian poxviruses). Differentiation of fowl, turkey, canary, and pigeon poxviruses on the basis of their pathogenicity for the host spectrum has been attempted. Although some strains show pathogenicity for several host species, others either are host-specific or infect a limited number of species (14, 15). Because natural pox infections have been reported in several species of wild birds of different families, as well as in domestic birds, it appears that all avian species are susceptible to avian pox. Minor antigenic differences in strains of fowl poxvirus can be detected by sodium dodecyl sulfate-polyacrylamide gel electrophoresis and immunoblotting (7, 9, 16, 24), although the majority of the antigens are cross-reactive. Antigenic profiles of quail poxvirus and isolates from Hawaiian forest birds are remarkably different from fowl poxvirus. Comparison of the genomic profile of fowl, pigeon, and junco poxviruses after digestion of their DNA with restriction enzymes and agarose gel electrophoresis revealed that the majority of DNA fragments co-migrate. However, most isolates could be distinguished by the presence or absence of one or more DNA fragments (9, 16, 24). On the other hand, genomic profiles of quail, canary, mynah and Hawaiian bird poxviruses are different from profiles of fowl poxvirus (7, 9, 28).

DIFFERENTIATION FROM CLOSELY RELATED AGENTS

The diphtheritic form of pox in birds with associated respiratory signs must be differentiated from other respiratory infections, especially infectious laryngotracheitis, an infection caused by a herpesvirus that produces intranuclear inclusions (5). Oral lesions caused by vitamin A, pantothenic acid and biotin deficiency in young chicks or by T-2 toxin may be mistaken for pox lesions. Diphtheritic pox lesions in doves and pigeons may be mistaken for lesions of Trichomonas gallinae, which are diagnosed by microscopic examination of smears or by culture (29). 118

Chapter 25

Pox

30. Tripathy, D. N., and W. M Reed. Pox. In: Diseases of Poultry, 11th Ed. Y. M. Saif, H. J. Bames, B. W. Calneck, J. R. Glisson, A. M Fadly, L. R. McDougald, and D. E. Swayne, Eds. Iowa State University Press, Ames, pp. 253-269, 2003. 31. Tulman, E. R., C. L. Afonso, Z. Lu, G. F. Kutish, and D. L. Rock. The genome of canarypox virus. J. Virol. 78:353-366,2004.

26. Tripathy, D. N., and L. E. Hanson. A smear technique for staining elementary bodies of fowlpox. Avian Dis., 20:609-610,1976. 27. Tripathy, D. N., L. E. Hanson, and W. L. Myers. Passive hemagglutination test with fowlpox virus. Avian Dis. 14:29-38,1970. 28. Tripathy, D. N., L. E. Hanson, and A. H. Killinger. Immunoperoxidase technique for detection of fowlpox antigen. Avian Dis. 17:274-278,1973. 29. Tripathy, D. N., W. M Schnitzlein, P. J. Morris, D. L. Janssen, J. K. Zuba, G. Massey, and C. T. Atkinson. Characterization of poxviruses from forest birds in Hawaii. J. Wildl. Dis. 36:225-230,2000.

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26 BUDGERIGAR FLEDGLING DISEASE AND OTHER AVIAN POLYOMAVIRUS INFECTIONS Branson W. Ritchie and Phil D. Lukert

SUMMARY. Avian polyomavirus, which belongs to the family Papovaviridae, genus Polyomavirus, causes acute, highly fatal disease in many young psittacines. Agent Identification. Detection of target segments of viral nucleic acid is most easily made by submitting tissues or swabs for polymerase chain reaction (PCR) based testing. Culture of the virus may be done in budgerigar embryo cell cultures initially, followed by culture in chicken embryo fibroblast cell cultures. Serologic Detection in the Host. The virus-neutralization test is the serologic test of choice. Confirmation of disease in birds with characteristic clinical or gross lesions requires demonstration of viral nucleic acid by in situ hybridization, immunohistochemical staining for viral proteins or documentation of viral particles by electron microscopy. Birds recovering from the chronic form appear normal, although some may die months later from kidney failure (4).

INTRODUCTION Avian polyomavirus (family Papovaviridae, genus Polyomavirus) causes an acute, highly fatal disease in budgerigar fledglings as well as peracute, acute, or chronic infections of many other species of psittacine birds. Disease is most common in young birds but has also been documented in adults. In budgerigars the mortality rate can vary from 25 to 100%. Substantial evidence exists that the virus is eggtransmitted, because intranuclear inclusion bodies are observed in 1day-old fledglings. All psittacines should be considered susceptible to natural infection, but some species develop clinical disease more readily than others. Passeriformes and some species of Falconiformes are also affected by avian polyomavirus. Gallinaceous birds are susceptible to natural infection by avian polyomavirus, but clinical disease has only been described experimentally in severely immunocompromised animals. Antibodies generated against psittacine polyomavirus cross-react with the polyomavirus of finches; however, nucleic acid sequences of these viruses differ. Avian polyomavirus differs from mammalian polyomaviruses in pathogenicity. Polyomaviruses of mammals typically produce disease only in immunocompromised hosts. CLINICAL DISEASE Budgerigars Acute fatal infections usually occur in fledglings from 1 to 15 days of age. Affected fledglings may die suddenly with no premonitory signs or they may exhibit abdominal distention, subcutaneous hemorrhages, and reduced feather formation. Some birds develop neurologic signs with ataxia and tremors of the head and neck several days before death. The death rate is highest in birds affected at an age of less than 15 days. After 15 days, birds that develop clinical signs may survive and develop feather abnormalities. Avian polyomavirus is considered to be one cause of French molt in budgerigars (1,3). Nonbudgerigar Psittacine Birds Both young and adult nonbudgerigar psittacine birds can experience clinical infections with avian polyomavirus. Although most infections are subclinical, the disease can be severe and often fatal in young birds. The peracute form with no premonitory signs is most common in young birds and as the age increases the acute form is usually observed. The signs include depression, anorexia, weight loss, diarrhea, subcutaneous hemorrhages, and dehydration. Birds normally die within 12^18 hr after developing clinical signs. Posterior paresis and paralysis within 18 hr of the onset of the disease is suggestive of polyomavirus infections. Eclectus parrots and caiques appear to be highly susceptible to disease (4). In addition to the peracute and acute forms a chronic/progressive disease has been described and is typified by weight loss, polyuria, poor feather formation, and recurring bacterial or fungal infections.

SAMPLE COLLECTION

For virus isolation a pool of liver, kidney, spleen, and heart tissues should be collected and submitted frozen in a plastic bag. Submit tissue from dying or recently dead birds. Serum samples for serology may be collected in capillary tubes, centrifuged to separate the erythrocytes, and stored or shipped under refrigeration (4). For the detection of viral DNA by PCR-based testing, cloacal swabs from live birds or swabs of tissue sampled at necropsy are preferred. Swabs used to culture bacteria are adequate and should be encased in a protective holder. Swabs used for anaerobic bacterial culture with gel-type transport media may interfere with the assay and should not be used (4). PREFERRED CULTURE MEDIA AND SUBSTRATES

The method of choice for isolation of virus is described by Bozeman et al (2). A 10% homogenized tissue suspension in cell culture medium containing antibiotics is prepared, and a monolayer of budgerigar embryo fibroblast (BEF) cells is inoculated. After several passages in BEF cells at 4- or 5-day intervals, most isolates will replicate in chick embryo fibroblast (CEF) cell cultures. The BEF cell cultures are prepared from 10 to-16-day-old budgerigar embryos. The medium used is 60% Ham’s F-10 with glutamine and 40% M-199 with Hanks’ salts with glutamine. This medium is enriched with 4% tryptose phosphate broth and 5% fetal bovine serum for growth. Maintenance medium has the fetal bovine serum reduced to 2%. AGENT IDENTIFICATION

Cell Culture Properties Cells with enlarged nuclei are first seen 48 hr postinfection with avian polyomavirus. By 72-96 hr, the cells begin to round up and are swollen, with greatly enlarged nuclei. Cells fixed and stained with Giemsa stain exhibit large, clear intranuclear inclusion bodies with margination of the chromatin. Infectivity titrations of the virus using cytopathic effect (CPE) as the endpoint are held for 7-9 days. Titration times may be reduced to 72 hr if immunofluorescence is used to detect viral infection.

Polymerase Chain Reaction A DNA probe based test has been developed and is commercially available (Infectious Diseases Laboratory, Athens, GA.). The DNA probe based test has been shown to be sensitive and specific (1). The sample of choice for detecting birds that are shedding viral nucleic acid is a cloacal swab. The same test can be used to detect viral nucleic acid in whole unclotted blood. A swab of the cut surface of the spleen, liver, and kidney can be submitted for PCR-based testing 120

Chapter 26 Budgerigar Fledgling Disease and Other Avian Polyomavirus Infections

to document the presence of viral nucleic acid in birds that have died with suspicious clinical and gross lesions. Physiochemical Properties Virions have a buoyant density of 1.34 g/cm3 in CsCl and a mean diameter of 42 mm (2). The virions are nonenveloped and contain a 4981-bp double-stranded DNA genome. Growth in cell culture is inhibited by 5-iododeoxyuridine. Freezing and thawing of the virus for five cycles has no effect on infectivity. The virus is relatively heat resistant. Incubation of the virus at 56 C for 30 min reduces the titer by 0.6 log10, and 56 C for 90 min reduces the titer by 1.0 logi0.

SEROLOGIC DETECTION IN THE HOST A virus-neutralization (VN) test is used for the detection of antibodies to avian polyomavirus. The test is performed in 96-well microtiter plates. Virus adapted to CEF cells is used in the test. A 100 mean tissue-culture infective dose (TCID50) dose of virus is used and twofold dilutions of serum added. The virus-serum mixture is incubated 30-60 min at 37 C and then a suspension of CEF cells is added in growth medium. After 72 hr the endpoint may be read by staining with a fluorescein-isothiocyanate-labeled anti-polyoma virus globulin or the cells can be held for 7-9 days to read the endpoint by CPE. Serum endpoints are determined as the last dilution of serum that prevents detectable infection of the cells. Serum titers of 1:20 or greater are considered to be positive for VN antibodies. No evidence of varying serotypes has been shown with avian polyomaviruses of psittacine birds (5).

DIFFERENTIATION FROM CLOSELY RELATED AGENTS

Because of the feathering abnormalities in chronic infections, polyomavirus infection may be confused with psittacine beak and feather disease. French molt can be caused by psittacine beak and feather disease virus, polyomavirus, or both. However, no beak lesions have been associated with polyomavirus infections. In addition, feather loss is not as extensive in polyomavirus infections as it is in psittacine beak and feather disease. Inclusion bodies associated with both viruses are different; those of polyomavirus are large, clear intranuclear inclusions and those of beak and feather disease virus are more basophilic and occur in the nucleus and cytoplasm. In situ hybridization can also be used to differentiate these viruses. REFERENCES

1. Bernier, G., M Morin, and G. Marsolais. Papovavirus induced feather abnormalities and skin lesions in the budgerigar: clinical and pathological findings. Can. Vet. J. 25:307-310. 1984. 2. Bozeman, L. H., R. B. Davis, D. Gaudry, P. D. Lukert, O. J. Fletcher, and M J. Dykstra. Characterization of a papovavirus isolated from fledgling budgerigars. Avian Dis. 25:972-980. 1981. 3. Davis, R. B., L. H. Bozeman, D. Gaudry, O. J. Fletcher, P. D. Lukert, and M J. Dykstra. A viral disease of fledgling budgerigars. Avian Dis. 25:179183. 1981. 4. Ritchie, B. W. Papovaviaridae. In. Avian Viruses: Function and Control, 1st ed. Wingers Publishing, Inc., Lake Worth, Fla. pp. 127-170. 1995. 5. Wainwright, P. Ο., P. D. Lukert, R. B. Davis, and P. Villegas. Serological evaluation of some Psittaformes for budgerigar fledgling disease virus. Avian Dis. 31:673-676. 1987.

27 PSITTACINE BEAK AND FEATHER DISEASE Branson W. Ritchie and Phil D. Lukert

SUMMARY. Psittacine beak and feather disease can be an acute or chronic disease of psittacine birds and is characterized by necrotic and abnormally formed feathers, beak necrosis and fractures, and usually death. The disease is caused by a small virus in the family Circoviridae. Agent Identification. The virus has not been cultured in vitro, but a DNA probe based test can be used to detect the target segments of viral nucleic acid in infected birds. Virus purified from infected birds will agglutinate cockatoo erythrocytes. Confirmation of disease in birds with characteristic clinical changes requires demonstration of viral nucleic acid by in situ hybridization, immunohistochemical staining for viral proteins or documentation of viral particles by electron microscopy. Serologic Detection in the Host. A hemagglutination-inhibition test can be used to detect antibodies to the virus, but it is of minimal value in managing the disease in individual birds or in aviaries.

INTRODUCTION

SAMPLE COLLECTION

Psittacine beak and feather disease (PBFD) is usually a chronic disease of psittacine birds that eventually leads to death. It was first described in various species of Australian cockatoos in the early 1970s (4). The virus that causes this disease belongs to the family of viruses called Circoviridae. The family includes viruses that infect Psittaciformes, Columbiformes, Passeriformes, and Anseriformes and are the smallest known virus of animals with a 14-17 nm diameter and circular single stranded DNA containing about 17002300 bases. In comparison, parvoviruses are 25 nm in diameter and have a single- stranded linear DNA with 5000 bases. Psittacine circovirus has not been grown in vitro, but it has been characterized by purifying the virus from the skin (feather follicles) of affected birds (2). All psittacine birds are probably susceptible to the virus and the outcome of infection is largely dependent upon the age of the bird when infected. Young birds are more susceptible to disease after infection. Generally, the older the bird the longer the incubation period between infection and appearance of disease. The disease usually occurs in birds less than 3 yr of age but birds up to 20 yr of age have been diagnosed with PBFD (1).

Samples that are useful for the detection of psittacine circovirus nucleic acid are white blood cells and swabs from feather pulp, necropsy tissue, or environmental areas. Blood samples should be collected in heparin (0.01 ml heparin to 0.49 ml of blood). Ethylenediaminetetraacetic acid or excessive heparin may result in false negative polymerase chain reaction (PCR) based tests. Clotted blood is unsuitable for nucleic acid detection. Swabs should be encased in a swab holder and anaerobic swabs with gel-type protective media are unacceptable. Serum samples for antibody detection by hemagglutination-inhibition (HI) tests may be submitted in microhematocrit capillary tubes after centrifugation to separate cells from the serum (1).

The virus has not been propagated in embryos or cell cultures, but can be identified and characterized by purifying the virus from homogenized preparations of skin from infected birds. The nucleic acid can be extracted and identified by gel electrophoresis or the virus visualized by electron microscopy.

CLINICAL DISEASE

AGENT IDENTIFICATION

The incubation period for the disease is highly variable and may be influenced by maternal antibodies, virus dose, or the host response to the infection. The minimum incubation period is 3-4 wk. Virus is thought to spread by direct contact or through contaminated food and water. Evidence exists that the virus can be transmitted vertically through the egg (1). The first clinical sign of disease is the appearance of necrotic or abnormally formed feathers. In young birds (less than 2 mo old), all of the feather tracts may be involved, whereas in older birds the disease is more prolonged with progressive feather changes occurring with ensuing molts. Peracute infections occur in neonates with signs of septicemia, pneumonia, enteritis, rapid weight loss, and death. The acute form is common in young birds during their first feather formation. Usually several days of depression occur, followed by sudden changes in developing feathers, which include necrotic, broken, bent, and hemorrhagic feathers. Crop stasis and diarrhea followed by death in 1-2 wk is a usual occurrence in the acute form of the disease. The chronic form of PBFD is characterized by symmetrical, progressive appearance of abnormal feathers during successive molts. The course of the disease may be protracted and the birds may experience secondary infections during this period. Birds with varying degrees of feathering may live months to years before eventual death, typically from a secondary infection. The beak lesions usually follow the feather malformations and changes include longitudinal fractures and necrosis of the palate area (1).

Polymerase Chain Reaction (PCR) A DNA probe based test has been developed and is available commercially (Infectious Diseases Laboratory, Athens, GA). The test is sensitive and specific (1). Most infected birds clear an infection with no signs of disease. Birds with no feather abnormalities that or blood positive for viral nucleic acid should be retested in 90 days. A negative test in the absence of clinical changes suggests that the bird has cleared the infection. A non-lory psittacine bird with a positive test and developmental feather lesions should be considered infected but disease can only be confirmed by histopathology. Lories have been shown to be susceptible to a variant of psittacine circovirus and birds with advanced feather abnormalities can recovery. Specific PCR-based testing is necessary to differentiate the two types of psittacine circovirus that may infect lories and is also required to provide accurate prognostic information. Once a psittacine aviary is free of PBFD virus a negative flock can be maintained through quarantine and testing procedures. Until a commercial vaccine is available, PCR-based testing will remain the most valuable tool for the control of this disease. In situ hybridization can be used to confirm a diagnosis of PBFD and differentiate it from budgerigar fledgling disease, which is caused by avian polyomavirus infections (1).

PREFERRED CULTURE MEDIA AND SUBSTRATES

Physicochemical Properties Virions are 14-17 nm in diameter and are nonenveloped. The viral DNA is single-stranded and circular, containing approximately 1800 bases. The virus has a density of 1.37 g/cm3 in CsCl (2), and is highly resistant to heat and disinfectants. Chicken anemia virus, 122

Chapter 27 Psittacine Beak and Feather Disease

which is in the same family (Circoviridae), resists 80 C for 1 hr and resists 5% phenol (5).

SEROLOGIC DETECTION IN THE HOST The virus will hemagglutinate erythrocytes of cockatoos but not those of sheep or chickens. An HI test can detect antibodies in infected birds and is typically used for epizootiological surveys. The HI test is, however, of minimal value in managing the disease in individual birds or in aviaries. Virus for the HI test is prepared by homogenizing feather follicle tracts from affected birds. The virus is concentrated on sucrose gradients followed by purification on CsCl gradients (3). DIFFERENTIATION FROM CLOSELY RELATED AGENTS Initially the chronic forms of PBFD may be confused with avian polyomavirus (budgerigar fledgling disease). The differentiation of the two diseases may be readily made by PCR-based testing of white blood cells from the bird. In some cases the intranuclear inclusions of avian polyomavirus can be differentiated from those induced by

PBFD virus. The PBFD inclusions tend to be basophilic and are found in the nucleus and cytoplasm, whereas polyomavirus produces large clear intranuclear inclusions. In situ hybridization with viral specific probes can be used to confirm a diagnosis (1). REFERENCES

1. Ritchie, B. W. Circoviridae. In: Avian Viruses: Function and Control, 1st ed Wingers Publishing, Inc., Lake Worth, Fla. pp. 223-252. 1995. 2. Ritchie, B. W., F. Niagro, P. Lukert, W. Steffans and K. Latimer. Characterization of a new virus from cockatoos with psittacine beak and feather disease. Virology 171:83-88. 1989. 3. Ritchie, B. W., F. Niagro, K. Latimer, P.Lukert, and D. Pesti. Hemagglutination by psittacine beak and feather disease virus and use of hemagglutination-inhibition for detection of PBFD virus antibodies. Am J. Vet. Res. 52:1810-1815. 1991. 4. Perry, R. A. A psittacine combined beak and feather disease syndrome. In: Proceedings # 55 of Courses for Veterinarians, Sydney, Australia, pp. 81108. 1981. 5. Yuasa, N. Effect of chemicals on the infectivity of chicken anemia virus. Avian Pathol. 21:315-319. 1992.

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28 CHICKEN ANEMIA VIRUS M. Stewart McNulty and Daniel Todd SUMMARY. Chicken anemia virus (CAV) causes a disease in 2 to 4 wk-old chickens characterized by increased mortality, anemia, thymic atrophy, and subcutaneous hemorrhages. Disease occurs following vertical transmission of CAV from a breeder flock. Agent Identification. Chicken anemia virus can be detected in affected chickens by virus isolation, immunocytochemical and immunofluorescence techniques, and a variety of molecular methods that detect CAV DNA. Serologic Detection in the Host. Enzyme-linked immunosorbent assay is the preferred method for detecting antibodies to CAV.

disease outbreaks have been made from liver, but the virus has also been isolated from skin, spleen, thymus, lung, heart, bursa, muscle, feces, and bone marrow. For virus isolation in susceptible chicks or in MDCC-MSB1 cells (a Marek’s disease virus transformed chicken lymphoblastoid cell line), 20% tissue suspensions are made using a stomacher or pestle and mortar in RPMI 1640 medium (GIBCO-BRL Laboratories, Gaithersburg, Md.) containing 1000IU penicillin per milliliter and 1000pg streptomycin per milliliter. Specimens are clarified by centrifugation at 2500 x g for 30 min before inoculation. Heat (70 C for 5 min) and chloroform treatment of inocula, using standard methods, may be performed before inoculation to inactivate other viruses that may be present. For detection of CAV antigens by immunofluorenscence, acetonefixed impression smears of thymus and bone marrow, and cryostat sections of thymus are prepared using standard methods (10). For immunocytochemistry, tissues should be fixed in 10% neutral buffered formalin for no longer than 6 hr; after this time destruction of CAV antigens occurs (1,24). Fixation for up to 7 days in neutral buffered formalin has no apparent effect on detection of CAV DNA in thymus by in situ hybridization (1).

INTRODUCTION Chicken anemia virus (CAV), formerly known as chicken anemia agent, is a small DNA virus that causes a disease in chickens called blue wing, infectious anemia, hemorrhagic syndrome, or anemia dermatitis syndrome. This disease is characterized by aplastic anemia, lymphoid depletion in the thymus, atrophy of hematopoietic tissues, subcutaneous and intramuscular hemorrhages, and increased mortality (11). CLINICAL DISEASE

Serological surveys have shown that most commercial chicken flocks (13) and many specific-pathogen-free (SPF) flocks (14) possess antibody to CAV. Outbreaks of clinical disease are rare. Onset of disease is acute and is usually seen from 2 to 4 wk of age. Disease occurs following infection of in-lay breeding flocks that have no antibody to CAV. No clinical signs are seen in the breeders, but CAV is vertically transmitted to the progeny. Affected chicks are produced from eggs laid over a period of 3-6 wk following infection. Diseased birds are depressed and weak, and they may be stunted. Mortality rate is variable, usually about 10%, but it may be as high at 60%. At necropsy, carcasses may be pale with thin, watery blood and yellowish bone marrow. Thymuses, and, to a lesser extent, bursas, are atrophic. Livers may be enlarged and yellowish. Skin lesions, in the form of ecchymotic hemorrhages, commonly occur on the wings. These lesions are prone to secondary bacterial infection leading to gangrenous dermatitis. Histologically, lymphocytes are depleted in the thymus, particularly in the thymic cortex. The bone marrow is aplastic, with hematopoietic tissue replaced by adipose tissue. The disease is usually seen in broilers, but it has been seen in replacement laying pullets. A similar condition in older chicks, sometimes accompanied by inclusion-body hepatitis, may be due to mixed infections of CAV and adenovirus. Vertically infected chicks excrete virus, resulting in horizontal spread to their hatchmates. Horizontally acquired CAV infection has been associated with impaired economic performance in broilers (16). The severity of the disease in horizontally infected chicks is variable, depending on factors that include age, virus dose and concurrent infections (12). To our knowledge, there are no peer-reviewed reports documenting the presence of CAV in avian species other than domestic fowl.

PREFERRED CULTURE MEDIA AND SUBSTRATES

Chicken anemia virus can be isolated either in cultured cells including MDCC-MSB1 and MDCC-CU147 (5) or by inoculation of susceptible chicks. Chick inoculation is more straightforward but requires the availability of chicks derived from SPF flocks that are free of CAV infection. Many SPF chicken flocks have antibody to CAV. Chicks used for isolation of CAV must be free of such antibodies, as the presence of maternal antibody in chicks confers resistance to experimentally induced disease caused by CAV. Although MDCC-CU147 cells appear to be susceptible to a wider range of CAV isolates and produce higher virus yields, MDCCMSB1 cells are the most commonly used cells for isolating and growing CAV. MDCC-MSB1 cells are grown in suspension in RPMI 1640 medium (GIBCO-BRL) with 5%-10% fetal bovine serum at 39 C in a 5% CO2 atmosphere. Following seeding at 2 x 105 to 5 x 105 cells per milliliter, cell cultures must be subcultured every 2-3 days. Following infection of MDCC-MSB1 cells with CAV, a cytopathic effect develops that is characterized by an increase in cell size, followed by lysis. CAV-infected MDCCMSB1 cell cultures are not capable of successful subculture. Cell death is also indicated by alkalinity of the medium. Two tubes or 2 wells in 24 well cell culture plates (Costar, Coming), each containing 1 ml MDCC-MSB1 cells seeded at 3 x 105 cells per milliliter, are each inoculated with 0.1 ml tissue suspension. Cultures are incubated at 39 C in a 5% CO2 atmosphere for 48 hr. If the inoculum appears to be toxic, subculture is carried out for the first two subcultures by transferring 0.3 ml medium and resuspended cells from inoculated cultures to 1 ml fresh cultures containing 3 χ 105 cells. Where toxicity is not a problem, subcultures are carried out by transferring 0.1-0.2 ml medium and cells from inoculated cultures to 1 ml warm growth medium every 48-72 hr. Isolation of CAV in MDCC-MSB1 cells may require up to 10 subcultures or passages before a cytopathic effect is evident.

SAMPLE COLLECTION

The easiest way to detect infected flocks is to examine sera for antibodies to CAV. Dead or sick chicks can be submitted for attempted virus isolation, detection of viral antigen, or detection of viral nucleic acid. Tissue samples for virus isolation can be sent to a laboratory in any virus transport medium. Although CAV is an extremely heat-resistant virus, samples should be kept cool during transport to discourage the growth of bacteria whose presence may complicate isolation of the virus. Most of the isolates of CAV from

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To isolate CAV in chicks, 1-day-old SPF chicks devoid of maternal antibody to CAV are inoculated intramuscularly with 0.1 ml tissue suspension. After 14 days, heparinized blood samples are taken, and the hematocrit levels are determined. Levels below 27% indicate anemia and suggest the presence of CAV. Following euthanasia, inoculated birds are examined grossly and histologically for CAV lesions. Because low doses of CAV do not produce anemia experimentally, some birds should be retained and bled again from 21 to 28 days postinoculation and examined for the presence of antibodies to CAV. More than one passage in SPF chicks may be necessary to produce anemia. Once experimental anemia and associated pathology have been reproduced, isolation of the virus from livers of inoculated chicks in MDCC-MSB1 cells is relatively straightforward. Alternatively, CAV antigens and/or nucleic acids can be demonstrated in the tissues of submitted and/or inoculated chicks.

Chicken Anemia Vine

0.1 ml of the virus-serum mixtures are inoculated into each of four tubes, each containing 0.9 ml MDCC-MSB1 cell suspension containing 3 * 105 cells per ml. Appropriate virus, serum, and positive and negative controls also should be included. MDCCMSB1 cells are then subcultured at 2-3 day intervals into fresh medium as described above. If the isolate is CAV, it will be neutralized by the CAV antiserum and no cytopathic effect will be evident. Antisera are prepared by intramuscular inoculation of 4-wk-old SPF chicks with 1 ml of cell-free CAV. A booster intravenous inoculation (1 ml) is given 4 wk later, and the birds are completely bled out after a further 3 wk (13). Hyperimmune rabbit antisera (10) or mouse monoclonal antibodies (15) can also be used. More recently developed diagnostic methods are quicker, less laborious, and/or cheaper than virus isolation. CAV antigens in diseased birds can be easily and rapidly detected by immunofluorescent staining of impression smears of thymus and bone marrow, and of cryostat sections of thymus. Immunofluorescence is concentrated in the thymic cortex. Antigens can also be detected by immunocytochemical staining of formalinfixed, paraffin-embedded tissues; however, for optimal results treatment of the sections with protease XIV (Sigma Chemical Company, St. Louis, Mo.) and fixation with formalin for no more than 6 hr are necessary (1,24). Immunocytochemical staining is more technically demanding, slower, and more laborious than immunofluorescence, and nonspecific staining can occur, but preservation of tissue morphology is much superior. Chicken anemia virus nucleic acids in tissues from diseased birds and infected MDCC-MSB1 cells dan be detected by a number of techniques, including in situ hybridization (1), dot-blot hybridization (19,30) and polymerase chain reaction (PCR) amplification. Dot-blot hybridization methods, which can be completed in 2-4 days, have a limit of sensitivity of only 105-106 genome copies (1—10 pg CAV DNA) and are likely to be less sensitive than virus isolation. Greater speed and sensitivity can be achieved using PCR amplification methods, many variations of which have been developed. PCR methods that involve the visual detection of PCR product following agarose gel electrophoresis, ethidium bromide staining and exposure to UV light generally have sensitivity limits of 10!-103 genome copies (0.1 - 10 fg) (D. Todd, unpubl. obs.). Hybridization techniques to detect PCR product have been employed to enhance sensitivity and to demonstrate specificity (19,20,25,27,33) but their inclusion makes these PCR methods more labor-intensive and time-consuming. Nested PCR tests offer the greatest sensitivity and have been used to detect CAV DNA in embryonal tissues and egg shell membranes (18) and to demonstrate the presence of CAV DNA in blood cells and reproductive tissues from chickens with neutralizing antibodies (2,7). The most sensitive PCR methods are likely to have greater sensitivity than virus isolation. While a highly sensitive PCR test may be the method of choice in research investigations and when examining biologicals for evidence of contamination with CAV, caution must be exercised in unreservedly recommending such tests for routine diagnosis of CAV. Given that low levels of CAV DNA can persist in chickens for months after infection, detection of virus DNA by a highly sensitive PCR technique may be of questionable value to the diagnostician faced with clinical disease problems. The ability to quantify CAV DNA, made possible by applying competitive (34) or real time (9) PCR methods, may assist in determining the clinicopathological significance of CAV in the clinical problem, but these methods will not be available in many diagnostic laboratories.

AGENT IDENTIFICATION

Chicken anemia virus is a non-enveloped, icosahedral (T = 1) virus, approximately 25 nm in diameter, comprising 60 structural units arranged as 12 pentamers (6). The virus possesses a single­ stranded, circular DNA genome, approximately 2.3 kb in size. CAV has a density of 1.33-1.34 g/ml in CsCl (12). CAV is classified as the type species of the genus Gyrovirus of the family Circoviridae (28). Chicken anemia virus is a remarkably stable virus. It resists treatment with lipid solvents such as chloroform and ether. It survives heating at 70 C for 1 hr and is only partially inactivated by a range of commercial disinfectants, including invert soap, amphoteric soap, orthodichlorobenzene, iodine disinfectant, and sodium hypochlorite, and by formaldehyde fumigation (35). Available evidence suggests that only one serotype of CAV exists. The existence of a second CAV serotype was suggested when an antigenically distinct agent was shown to cause similar pathogenic effects following experimental infection of SPF chickens (26). However, the causal agent needs to be isolated and molecularly characterized to confirm whether it represents a second CAV serotype. Minor antigenic differences between CAV isolates have been revealed by monoclonal antibody binding studies (15). Naturally occurring isolates of CAV obtained from chickens appear to possess similar pathogenicity for young chicks. Naturally occurring apathogenic isolates have been obtained from turkeys (23). CAV can be attenuated by passage in MDCC-MSB1 cells (29) or chicken embryos (22). CAV does not agglutinate the commonly available avian and mammalian erythrocytes (D. Todd, unpubl. obs.). The earliest method to identify CAV was isolation of the virus in SPF chicks or MDCC-MSB1 cells, as described above. Confirmation of the identity of putative isolates is achieved by immunofluorescent staining of infected MDCC-MSB1 cells. Infected MDCC-MSB1 cells are pelleted by centrifugation at 1500 x g for 10 min, resuspended in a minimal volume of phosphate-buffered saline, and smeared onto multispot Tefloncoated slides. These are air-dried, fixed in acetone for 10 min at room temperature, and stained by direct or indirect immunofluorescence using chicken antisera to CAV or monoclonal antibodies that are known to react with all CAV isolates (15). CAV antigens are found in the nuclei of infected cells. Fine granular fluorescence usually is present throughout the nuclei, with some nuclei containing larger spherical inclusions. Isolates growing in MDCC-MSB1 cells also can be confirmed as CAV using a neutralization test. Chicken antiserum to CAV and control chicken serum without antibody to CAV are diluted 1:80 in RPM1 1640 containing antibiotics, inactivated by treatment at 56 C for 30 min, cooled, and then added to an equal volume of the undiluted isolate and incubated for 30 min at 37 C. Samples of

SEROLOGIC DETECTION IN THE HOST Antibodies to CAV were originally detected by indirect immunofluorescence and virus neutralization (4,12). However, both of these tests have now been superseded by enzyme-linked

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M Stewart McNulty and Daniel Todd 10. McNeilly, F., G. M Allan, D. A. Moffett, and M S. McNulty. Detection of chicken anaemia agent in chickens by immunofluorescence and immunoperoxidase staining. Avian Pathol. 20:125-132. 1991. 11. McNulty, M S. Chicken anemia agent. In: A laboratory manual for the isolation and identification of avian pathogens, 3rd ed. H. G. Purchase, L. H. Arp, C. H. Domermuth, and J. E. Pearson, eds. Kendall/Hunt Publishing Company, Dubuque, Iowa. pp. 108-109. 1989. 12. McNulty, M S. Chicken anaemia agent: a review. Avian Pathol. 20:187-203. 1991. 13. McNulty, M S., T. J. Connor, F. McNeilly, K. S. Kirkpatrick, and J. B. McFerran. A serological survey of domestic poultry in the United Kingdom for antibody to chicken anaemia agent. Avian Pathol. 17:315-324. 1988. 14. McNulty, M S., T. J. Connor, and F. McNeilly. A survey of specific­ pathogen-free chicken flocks for antibodies to chicken anaemia agent, avian nephritis virus, and group A rotavirus. Avian Pathol. 18:215-220. 1989. 15. McNulty, M S., D. P. Mackie, D. A. Pollock, J. McNair, D. Todd, K. Mawhinney, T. J. Connor, and F. McNeilly. Production and preliminary characterization of monoclonal antibodies to chicken anemia agent. Avian Dis. 34:352-358. 1990. 16. McNulty, M S., S. G. McIlroy, D. W. Bruce, and D. Todd. Economic effects of subclinical chicken anemia agent infection in broiler chickens. Avian Dis. 35:263-268. 1991. 17. Michalski, W. P., D. O’Rourke, and T. J. Bagust. Chicken anaemia virus antibody ELISA: problems with non-specific reactions. Avian Pathol. 25:245-254. 1996. 18. Miller, Μ Μ, K. A. Ealey, W. B. Oswald,, and K. A Schat. Detection of chcken anemia virus DNA in embryonal tissues and eggshell membranes. Avian Dis. 47: 662-671. 2003. 19. Notebom, Μ Η. M, C. A. J. Verscheuren, D. J. van Roozelaar, S. Veldkemp, A. J. van der Eb, and G. F. de Boer. Detection of chicken anaemia virus by DNA hybridization and polymerase chain reaction. Avian Pathol. 21:107-118. 1992. 20. Novak, R., and W. L. Ragland. Competitive DNA hybridization in microtitre plates for chicken anaemia virus. Mol. Cell. Probe. 15:1-11. 2001. 21. Pallister, J., K. J. Fahey, and M Sheppard. Cloning and sequencing of the chicken anaemia virus (CAV) ORF-3 gene, and the development of an ELISA for the detection of serum antibody to CAV. Vet. Microbiol. 39:167-178. 1994. 22. Schrier, C. C. Chicken anaemia agent vaccine. European Patent Application 92202864.2. 1992. 23. Schrier, C. C. and H. J. M Jagt. Chicken anaemia viruses of low pathogenicity. US Patent Application 2001/0023664 Al 24. Smyth, J. A, D. A. Moffett, M S. McNulty, D. Todd, and D. P. Mackie. A sequential histopathologic and immunocytochemical study of chicken anemia virus infection at one day of age. Avian Dis. 37:324-328. 1993. 25. Soine, C., S. K. Watson, E. Rybicki, B. Lucio, R. M Nordgren, C. R. Parrish, and K. A. Schat. Determination of the detection limit of the polymerase chain reaction for chicken infectious anemia virus. Avian Dis. 37:467^176. 1993. 26. Spackman, E., S. S. Cloud, C. R. Pope, and J.K. Rosenberger. Comparison of a putative second serotyoe of chicken infectious anemia virus with a prototypical isolate. I. Pathogenesis. Avian Dis. 46: 945-955. 2002. 27. Tham, K. M, and W. L. Stanislawek. Detection of chicken anaemia agent DNA sequences by the polymerase chain reaction. Arch. Virol. 127:245-255. 1992. 28. Todd, D., M Bendinelli, P. Biagini, S. Hino, A. Mankertz, S. Mishiro, C. Niel, H. Okamoto, S. Raidal, B. W. Ritchie, and G. C. Teo. Circoviridae. In: Virus Taxonomy, VUIth Report of the International Committee for the Taxonomy of Viruses. C.M Fauquet, MA. Mayo, J. Maniloff, U. Desselberger, and L.A. Ball, eds. , Elsevier/Academic Press, London, pp 327-334. 2004. 29. Todd, D., T. J. Connor, V. M Calvert, J. L. Creelan, B. M Meehan, and M S. McNulty. Molecular cloning of an attenuated chicken anaemia virus isolate following repeated cell culture passage. Avian Pathol. 24:171-187. 1995. 30. Todd, D., J. L. Creelan, and M S. McNulty. Dot blot hybridization assay for chicken anemia agent using a cloned DNA probe. J. Clin. Microbiol. 29:933-939. 1991. 31. Todd, D., D. P. Mackie, K. A. Mawhinney, T. J. Connor, F. McNeilly, and M S. McNulty. Development of an enzyme-linked immunosorbent assay to detect serum antibody to chicken anemia agent. Avian Dis. 34:359-363. 1990.

immunosorbent assay (ELISA). A number of ELISAs have been described (3,8,21,31,32). Most use CAV grown in MDCC-MSB1 cells as antigen. With the indirect ELISA format, antigen-coated plates are reacted with dilutions of chicken sera and the presence of chicken antibodies specific to CAV are detected using an anti­ chicken immunoglobulin enzyme conjugate. In the competitive or blocking format, the presence of virus-specific chicken antibodies in test sera is detected when the reaction between the coated antigen and an enzyme-conjugated virus-specific mouse monoclonal antibody is prevented or reduced. Competitive or blocking tests generally exhibit fewer problems in relation to the non-specific binding of chicken antibodies to cellular antigens. ELISAs are now available commercially (Flockscreen Chicken Infectious Anaemia Antibody ELISA kit, Guildhay Ltd. Guildford, Surrey, United Kingdom; Chicken Anemia Virus (CAV) ELISA test kit, IDEXX Laboratories, Inc., Westbrook, Maine). Lack of specificity with some ELISAs is not a major problem in serologic profiling of commercial flocks, but can cause problems when testing SPF flocks (17). Following an outbreak of anemia dermatitis syndrome, retrospective testing of sera from the breeder flocks, if sera are available, will identify one or more breeder flocks that were seronegative at point-of-lay, but that have developed antibodies by the time disease is recognized in their progeny. These breeder flocks are probably the source of vertically transmitted virus. DIFFERENTIATION FROM CLOSELY RELATED AGENTS

The differential diagnosis of anemia dermatitis syndrome includes those conditions causing anemia, immunosuppression, and increased mortality. REFERENCES

1. Allan, G. M, J. A. Smyth, D. Todd, and M S. McNulty. In situ hybridization for the detection of chicken anemia virus in formalin-fixed, paraffin-embedded sections. Avian Dis. 37:177-182. 1993. 2. Brentano, L., S. Lazzarin, S. S. Bassi, T. A. P. Klein, and K. A Schat. Detection of chicken anemia virus in the gonads and in the progeny of broiler breeder hens with high neutralizing antibody titres. Vet. Micro. 105: 65-72. 2005. 3. Brewer, J., J. M Saunders, and N. J. Chettle. The development of an enzyme linked immunosorbent assay to detect antibodies to chicken anaemia virus, and its comparison with the indirect immimofluorescent antibody test. In: Proceedings of the International Symposium on Infectious Bursal Disease and Chicken Infectious Anaemia, Rauischholzhausen, Germany, pp. 408-412. 1994. 4. Bulow, V. V., B. Fuchs, and M Bertram. Untersuchungen uber den Erreger der infectiiosen Anamie bei Huhnerkuken (CAA) in vitro: Vermehrung, Titration, Serumneutralisationtest and indirekter Immunofluoresenztest. Zentralbl. Veterinaermed. Reihe B 32:679-693. 1985. 5. Calnek, B. W., B. Lucio-Martinez, C. Cardona, R. W. Harris, K. A Schat, and C. Buscaglia. Comparative susceptibility of Marek’s disease cell lines to chicken infectious anemia virus. Avian Dis. 44:114-124. 2000. 6. Crowther, R. A, J. A. Berriman, W. L. Curran, G. M Alan, and D. Todd. Comparison of the structures of three circoviruses: Chicken anemia virus, Porcine circovirus type 2 and Beak and feather disease virus. J. Virol. 77: 13036-13041. 2003. 7. Imai, K., M Mase, K. Tsukamoto, H. Hihara, and N. Yuasa. Persistent infection with chicken anaemia virus and some effects of highly virulent infectious bursal disease virus infection on its persistency. Res Vet. Sci. 67: 233-238. 1999. 8. Lamichlane, C. M, D. B. Snyder, T. Girschick, M A. Goodwin, and S. L. Miller. Development and comparison of serologic methods for diagnosing chicken anemia virus infection. Avian Dis. 36:725-729. 1992. 9. Markowski-Grimsrud, C. J., Μ M Miller, and K. A. Schat. Development of a strain-specific real-time PCR and RT-PCR assays for quantitation of chicken anemia virus. J. Virol. Meth. 101: 135-147. 2002.

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Chicken Anemia Virus

34. Yamaguchi, S., N. Kaji, Η. M Munang’andu, C. Kojima, M Mase, and K. Tsukamoto. Quantification of chicken anaemia virus by competitive polymerase chain reaction. Avian Path. 29: 305-310. 2000. 35. Yuasa, N. Effect of chemicals on the infectivity of chicken anaemia virus. Avian Pathol. 21:315-319. 1992.

32. Todd, D., K. A. Mawhinney, D. A. Graham, and A. N. J. Scott. Development of a blocking enzyme-linked immunosorbent assay for the serological diagnosis of chicken anaemia virus. J. Virol. Meth. 82: 177-184. 1999. 33. Todd, D., K. A. Mawhinney, and M S. McNulty. Detection and differentiation of chicken anemia virus isolates by using the polymerase chain reaction. J. Clin. Microbiol. 30:1661-1666. 1992.

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29 AVIAN INFLUENZA David E. Swayne, Dennis A. Senne and David L. Suarez SUMMARY. Avian influenza (Al) is caused by type A influenza virus, in the family Orthomyxoviridae. Type A influenza viruses are serologically categorized into 16 hemagglutinin (H1-H16) and 9 neuraminidase (N1-N9) subtypes. All subtypes have been identified in birds. Infections by influenza virus have been reported in a variety of domestic and wild birds, but most infections are subclinical. In poultry, influenza virus infections can be clinical, but the features are variable depending on virus strain, host species and breed, host age, concurrent infections, physiologic stresses, and environmental factors. Some H5 and H7 strains are highly pathogenic for poultry, producing a systemic disease with high mortality, and all H5 and H7 strains should be considered to have the potential to mutate from low (LP) to high pathogenicity (HP). Both LP and HP H5 and H7 strains isolated from poultry are notifiable to the World Organization for Animal Health (OIE). Agent Detection or Identification. Although several diagnostic tests can be used for detection of avian influenza, the initial diagnosis is preferably made by virus isolation via inoculation of embryonating chicken eggs, demonstration of hemagglutinating activity, and verification as avian influenza virus by agar-gel immunodiffusion (AGID) or other antigen-detection tests. Once a virus has been isolated, additional biologic and molecular studies can be performed to characterize the virus that isn’t possible by other means. This includes virus subtyping, testing of pathogenicity in chickens or other species, and for H5 and H7 subtypes the determination of amino acid sequence at the hemagglutinin proteolytic cleavage site to develop the appropriate control strategies. We do have an increased ability to rapidly diagnose and characterize Al by detection of nucleic acids specific for Al viruses and potentially by rapid and sensitive immunoassays, but these tests will not replace virus isolation in our ability to characterize Al viruses. Serologic Detection in the Host. Serologic detection of infection in the host is important for moderate-to-large scale surveillance by industry or government and for early diagnostic detection programs. Detection of type A group-specific antibodies via AGID or enzymelinked immunosorbent assay is preferred as the initial step for detection of primary infection, but confirmation and subtype determination via hemagglutination-inhibition and neuraminidase-inhibition tests are crucial adjuncts.

INTRODUCTION

of the head, comb, and wattles; and hemorthages on the serosal and mucosal surfaces of viscera including the proventriculus, on the shanks of the legs, and on the comb and wattles. Ruptured mature ovarian follicles (yolks) and accompanying inflammation ("peritonitis”) are frequently observed in active layers with either LP or HPAl viral infections. Birds that die peracutely may exhibit few, if any, obvious clinical signs or gross lesions. The period of time during which virus may be recovered after infection depends greatly upon the strain of virus, the type of sample, and the host species and varies from 1-28 days in domestic poultiy (29) and probably longer for wild species. However, AIV is infrequently recovered from experimentally infected poultry after 14 days. Detailed reviews on avian influenza are available (2,23,28).

Avian influenza virus (AIV) infections occur in a variety of domestic and wild bird species. The endemic nature of AIV in wild birds including ducks, gulls and shorebirds provides for a constant but low risk of introduction into poultry populations. In North America, infections in commercial poultry occur primarily in domestic turkeys, with avian and swine influenza viruses. However, outbreaks can be seen in domestic and imported exotic birds, ratites, and occasionally in chickens, quail, and upland game birds. Infections in the latter three groups have been detected most often in live bird marketing systems traditionally found in large metropolitan settings. The greatest incidence of the disease in domestic poultry occurs where there is opportunity for co-mingling or indirect contact with waterfowl, seabirds, and/or pigs. Historically, the best-known avian influenza virus (AIV) is fowl plague virus, an H7 subtype, which has caused poultry losses since the late 1800s in many parts of the world. The term fowl plague has been replaced by highly pathogenic or high pathogenicity avian influenza (HPAl) and is caused by some H5 and H7 strains (5). Epizootics of HPAl since 1995 have included: 1) H7N3 in Pakistan during 1995, 2001 and 2004; 2) H5N1 in Asia during 1996-2005; 3) H7N4 in Australia in 1997; 4) H5N2 in Italy during 1997; 5) H7N1 in Italy during 1999-2000; 6) H7N3 in Chile during 2002; 7) H7N7 in The Netherlands during 2003; 8) H7N3 in Canada during 2004; 9) H5N2 in USA during 2004; 10) H5N2 in South Africa during 2004, and 11) H7N7 in North Korea (9,10,12,14,16,28). Where there is suspicion or confirmation of Al, state and/or federal authorities should be notified.

SAMPLE COLLECTION

For Virus Isolation Samples from respiratory (trachea, lung, air sac, and sinus exudate) and digestive systems are suitable for virus isolation. They can be swabs or tissue. Cloacal swabs are typically best for AIV isolation from wild birds or domestic ducks. However, both tracheal or oropharyngeal, and cloacal swabs should be collected from domestic poultry. Some HPAl viruses can be isolated from liver, spleen, blood, heart, or brain of clinically ill birds. Dry sterile swabs preferably of a synthetic material, such as Dacron and not calcium alginate, should be used to obtain samples of tracheal, oropharyngeal or sinus exudate and of cloacal contents. Swabs with wooden shafts should be avoided unless the swabs are immediately removed from the specimen tube and discarded after expelling the swab contents into the transport medium (the wooden shaft may be treated with formalin that can interfere with virus/RNA detection). The swabs are placed in 2-3.5 ml of sterile transport medium prepared from brain-heart infusion, tryptose, nutrient, or peptone broth, or cell-culture medium containing 1% bovine serum albumin. A soft copper or nichrome loop may be used for small birds (19). Specimens should be kept at 4 C after collection and during transport. Low levels of antibiotics, e.g. 200 pg/ml gentamicin sulfate and 5 pg/ml amphotericin B, can be added to the transport medium to reduce growth of bacterial and fungal contaminants during transportation but antibiotics should used as an

CLINICAL DISEASE

The outcome of infection produced by AIV isolates varies from no obvious clinical signs to 100% mortality. Birds of all ages and most, if not all, avian species are susceptible to infection. Common signs or lesions in the major poultry species (chickens and turkeys) infected with LPAI viruses include slight-to-severe declines in egg production; increased mortality; diarrhea; respiratory difficulties; and occasionally, deposits of urates in the kidneys. Infections with HPAl viruses results in severe disease which can present as edema

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adjunct to chilling and not as an alternative to chilling. When isolation cannot be attempted immediately, samples can be stored at 4 C for up to 48 hr, or frozen (preferably at -70 C) for longer periods. Samples can be shipped with frozen gel packs if they will arrive at the laboratory within 48 hr.

(20 gg/ml), gentamicin sulfate (1000 pg/ml), and kanamycin sulfate (650 pg/ml). Supernatant should be kept at 22-25 C for 30 to 60 min just before inoculation, to allow the antibiotics to reduce the level of bacterial contamination. If further reduction in bacterial contamination is required to reduce embryo deaths or non-viral hemagglutinating activity (HA) of egg fluids, the supernatant can be filtered through pre-wet 0.22-0.45-μιη reusable or disposable membrane filters. However, filtration can remove low levels of virus from samples and reduce isolation rates. For tissue specimens, a 10% ground suspension is made in antibiotic-containing brain-heart infusion, tryptose, nutrient, or peptone broth, or cell-culture medium containing 1% bovine serum albumin. Grinding is accomplished in a mechanical homogenizer, stomacher, or mincing with sterile scissors and ground either in a sterile glass tissue grinder or with a sterile mortar and pestle using sand or similar abrasive materials. Alternatively, small tissue fragments (1 cm3) are placed in 2-3.5 ml medium, frozen and thawed, vigorously vortexed, centrifuged, and the supernatant diluted in equal volumes of medium containing antibiotics. Tissue suspensions are clarified by low-speed centrifugation (1000-1500 x g) and placed at room temperature for 30 to 60 min before inoculating embryos. Chicken embryos, 9-11 days old, are injected via the allantoic sac (see Chapter 43 on virus propagation in embryonating eggs) with 0.2-0.3 ml of clinical specimen. However, inoculation of embryos via the yolk sac route or the use of cell cultures (MDCK or VERO) with added trypsin has been more sensitive for the isolation of swine-origin influenza viruses from poultry. Three to five embryos per sample are usually adequate. Eggs are incubated at 35-37 C for 3-5 days. If virus is present, some embryos may die following inoculation, but some may not, depending on the virulence of the virus. Amnioallantoic fluid (AAF) from eggs with dead embryos as well as from eggs with surviving embryos should be collected and analyzed for HA. A second passage may be required for some viruses. Additional passages can be made by diluting the AAF 1:10 in antibiotic medium and inoculating a new set of eggs. The eggs from additional passages are incubated and tested using the same procedure as for the first passage.

For Reverse Transcriptase Polymerase Chain Reaction (RTPCR) Specimens suitable for virus isolation are generally suitable for RT-PCR (see collection of samples for virus isolation above). However, tracheal or oropharyngeal swabs are the specimens of choice for gallinaceous birds because they are sensitive, easy to process, and generally contain less extraneous organic material that can interfere with RNA recovery and amplification by PCR. Caution should be used when testing cloacal swabs by real-time RT-PCR (RRT-PCR) because of the concerns with false negative results due to poor RNA recovery or presence of PCR interfering substances. The use of RT-PCR for screening of environmental samples needs to be done with caution because there may be a poor correlation between a positive PCR result and presence of viable virus; e.g. inactivated virus cannot be isolated but may yield a positive RRT-PCR test result. Therefore, RT-PCR is not recommended to assure an environment free of avian influenza virus, because of the potential for the test to identify inactivated virus, providing a false positive. Calcium alginate swabs should be avoided for collection of samples for RT-PCR because they are reported to produce inhibition for PCR. When using RRT-PCR, particular attention should be given to the selection of an appropriate method for RNA extraction to maximize recovery. The RNA extraction step is considered one of the key steps for a successful test. Several commercial kits are suitable for the recovery of RNA from clinical specimens. The isolation of good quality RNA that is void of PCR inhibiting substances is essential for accurate test results. Organic extraction methods such as Trizol (Invitrogen, Carlsbad, CA) are generally better than silica gel membrane kits for most specimens and are especially good for cloacal swabs and tissues that have a high organic content. However, the toxic chemicals (phenol, guanidine isothiocyanate, and chloroform), and additional labor and processing time required for this method makes it less desirable than other methods. Silica gel membrane kits such as RNeasy (Qiagen, Valencia, CA) have been successfully used for processing tracheal and oropharyngeal swabs, but this system has lower sensitivity for tissue suspensions and cloacal swabs. This is most likely because of the heavy organic load in these specimens produces extraneous RNA or other substances that compete for binding sites on the silica membrane resulting in lower recovery rates of target RNA. The MagMAX kit (Ambion, Austin, TX) uses specially engineered magnetic beads to absorb RNA and is more resistant to RNA saturation problems associated with silica gel membrane kits. The MagMAX kit can be performed in 96-well plates and therefore is more suitable than other extraction methods for high throughput testing of oropharyngeal or tracheal swabs. However, the suitability of this method for extraction of tissue suspensions and cloacal swabs has not yet been fully evaluated.

RT-PCR RT-PCR procedures have been developed to aid in the detection of ATV in clinical samples and to facilitate assessment of pathogenicity of H5 and H7 ATV (18,33). Both traditional RT-PCR assays with analysis of the PCR product on an agarose gel and real­ time RT-PCR (RRT-PCR) assays have been described in the literature. Most of the methods have levels of sensitivity comparable to virus isolation, and are being commonly used as primary screening tests for avian influenza throughout the world. Of the two commonly used methods, each has its strengths and weaknesses. The traditional RT-PCR requires less expensive equipment that is available in many diagnostic laboratories, but does require additional labor and cross contamination is an ongoing concern. The RRT-PCR tests are less labor intensive, can be performed more rapidly, and have a reduced probability of cross contamination in the laboratory, but it does require expensive equipment to perform. Although many different RT-PCR tests for avian influenza have been described, few have been extensively validated for fitness for purpose with field samples in comparison to the performance standard of virus isolation. In general, both RT-PCR assays have four steps: 1) isolation of the RNA; 2) conversion of the viral RNA to cDNA by the use of reverse transcriptase; 3) amplification of the cDNA by PCR; and 4) evaluation of the PCR product (amplicon) for the presence or absence of the target DNA sequence. For traditional RT-PCR the specificity of the amplicon is confirmed by electrophoresis of the amplicon in an agarose gel containing ethidium bromide, then comparing the size of the amplicon to markers of known molecular

PREFERRED CULTURE MEDIA AND SUBSTRATES

Virus Isolation The preferred method of diagnosis requires isolation of AIV from clinical swabs or tissue in embryonating chicken eggs. For cloacal, tracheal, oropharyngeal, or sinus swabs received in appropriate transport media, the fluid is centrifuged at low speed (1000-1500 x g) to sediment debris and the supernatant is added to antibiotic medium. The following are suggested antibiotics and the maximum dose per ml that can be used singularly or in combination: penicillin (10,000 IU/ml), streptomycin (10,000 pg/ml), amphotericin B 129

David E. Swayne, Dennis A. Senne and David L. Suarez

weight. The detection of the PCR product of the proper size however provides only a presumption of the specific product, and additional confirmation can be provided by several methods including enzyme restriction enzyme endonuclease assays, Southern Blot hybridization, or by direct sequencing. The need for post­ amplification processing of the amplicon is one of the major drawbacks of using traditional RT-PCR as a diagnostic assay because of the increased labor and the high potential for contamination of the laboratory and cross contamination of samples. Extreme precautions, such as the use of dedicated equipment and separation of sample processing activities from post amplification processing activities, must be taken to prevent cross contamination. The RRT-PCR assays differ primarily in the analysis of the amplicon, which is done with fluorescently labeled probes or dyes and special thermocyclers that accurately measure light levels at different wavelengths. Several different strategies are available for the fluorescently labeled probes, including Taqman probes, Scorpion probes, Fret probes and others. The Taqman probes, also known as hydrolysis probes, are the most commonly used, and includes both an excitation dye and a quencher dye. Normally, the light produced from the excitation dye is dampened by the close proximity of the quencher dye. However, when the Taqman probe binds to the PCR product, the excitation dye is cleaved away from the probe and quencher dye by Taq polymerase, and the light produced from the excitation dye is no longer dampened by the quencher dye. This light produced is quantitative and is proportional to the amount of PCR product present in the reaction. The light level is taken after every PCR cycle, and the results can be visualized in real-time with no additional post-PCR processing. Because the probe must specifically bind to the PCR product, an additional level of assurance is provided that the PCR is specific for the intended agent. Although many different RT-PCR tests and even alternative RNA amplification tests have been described, most of these have never been validated for use with field samples. Validation is the determination of fitness for a particular use, and includes testing of a prescribed number of positive and negative field samples against a performance standard. In the U.S. a RRT-PCR assay described by Spackman et al. for detection of ATV RNA in clinical specimens from poultry was developed and validated with clinical specimens (n = 1,550) from live-bird market surveillance (22). The AIV assay was further validated (n = 3,500) during an outbreak of LPAIH7N2 in Virginia in 2002. The assay has three sets of primers and probes that are used in separate reactions, one to identify any type A influenza viruses (Matrix assay), one to identify H5 strains and one to identify H7 strains of ATV. The matrix test is targeted to a highly conserved region of the influenza genome, and it can detect any type A influenza virus, including swine, equine, and human influenza viruses. Because the hemagglutinin gene is known for its high variability, it has been difficult to develop a single test that would identify all H5 or all H7 viruses. The test validated for use in the U.S. has been shown to identify most H5 viruses from around the world, but the H7 test is more restricted in its geographic reach. The test used in the U.S. will identify North American origin H7 viruses, but it will not detect Eurasian H7s. Alternative primer/probe sets are available for H7s from other geographic regions, and the diagnostician must be aware of the strains that are likely to occur in their region. Since the matrix (M) assay has the highest level of sensitivity, samples are first screened with the M primers/probe then positive specimens are tested with the H5 and H7 assays. In the U.S. this assay is considered an official test for avian influenza, and it being used by more than 39 state laboratories, that have demonstrated proficiency in testing, as a front line surveillance tool for Al. The use of rapid molecular diagnostic tools can still be considered to be in its early stages, and additional technical advances should be expected. These incremental improvements in performance are

likely to come by improvements in the critical control points of RNA extraction, RT-PCR amplification, and primer and probe design. Opportunities to increase sensitivity, specificity, throughput and ease of use are likely to occur, and will become adopted as part of the official protocol. Additionally, consideration for ongoing analysis of test performance needs to be considered for all molecular diagnostic tests, since specificity is directly related to primer and probe design, and genetic variation or drift may affect test performance. Future research goals will include the use of an internal positive control (unrelated to the target cDNA) is also highly recommended to monitor for presence of PCR inhibiting substances that could cause a false negative test results. Several different strategies for an internal control have been described.

AGENT IDENTIFICATION Physicochemical, Morphologic and Biological Properties The influenza viruses are a diverse group of viruses that belong to the family Orthomyxoviridae, are enveloped (ether-sensitive), and contain single-stranded ribonucleic acid (RNA) that is segmented and has negative polarity. Two major internal components, the matrix protein (M) and ribonucleoprotein (RNP), are group-specific proteins that designate type specificity (i.e., A, B, or C). There are two surface glycoproteins, the hemagglutinin (H) and neuraminidase (N) glycoproteins, that project from the lipid membrane and participate in the infection process and define subtype specificity (i.e., H1-H16 and N1-N9). Influenza virions can be either spherical forms or short rods (80-120 nm) or filamentous forms (400-800 nm long).· Influenza viruses are relatively stable at pH 7-8 but are labile at the low pH range. Al viruses are sensitive to heat inactivation with inactivation at pasteurization temperatures (55.6-63.3 C) in 3 to 6.2 min or minimal cooking temperature (70 C) in a few seconds (26,27).

Initial Identification Hemagglutinating Activity (HA). The surface H glycoprotein of ATV isolates will bind to receptors on a variety of mammalian and avian erythrocytes, and this phenomenon is the basis for screening of AAF for presence of hemagglutinating agents. Small amounts of AAF can be harvested aseptically from eggs 24—48 hr after inoculation with a syringe and needle without killing the embryo (see Chapter 44 on virus propagating in embryonating eggs). This early sampling can speed diagnosis because HA or specific ATV antigen can be detected long before the embryo dies. HA is determined by making twofold dilutions in a microtiter plate followed by the addition of an equal volume of 0.5% washed chicken erythrocytes. In some cases alternative RBCs need to be considered. In particular the Hl and H3 swine viruses that commonly infect turkeys no longer hemagglutinate chicken RBCs. The use of turkey RBCs should be considered as an alternative, although guinea pig and horse RBCs may be used (25,30). Diluting the AAF avoids the occasional prozone effect that can occur with undiluted egg fluids. If HA is observed, the remaining AAF should be harvested after the egg has been chilled for 4—12 hr at 4 C. Chilling kills the embiyo and lessens the chance of contaminating the fluids with erythrocytes, which can absorb influenza virions. Embryos that survive the incubation period and AAF that lacks HA can be discarded after a sample of fluid is removed for an additional egg passage. Fluids from AJV-infected eggs occasionally may fail to exhibit HA, particularly when virus levels are less than that needed to cause hemagglutination, usually ΙΟ5—106 mean embryo infectious dose (EID5o)/ml. If the fluids are HA-negative, an additional passage is optional to be certain that isolations are not missed because of low levels of virus in the sample.

130

Chapter 29 Avian influenza

After demonstrating HA in AAF, the agent responsible for such HA must be determined. Common hemagglutinating agents for birds include AIV, avian paramyxoviruses including Newcastle disease virus (NDV) (see Chapter 30), a hemagglutinating adenovirus (see Chapter 19), and hemagglutinating bacteria. For the initial determination, some diagnostic laboratories run microtiter hemagglutination-inhibition (HI) tests to ascertain the presence or absence of NDV. If NDV antiserum inhibits the HA, NDV is present, but this inhibition does not exclude the possibility that the fluid also contains one or more other viruses. Confirmation of AIV in HA-positive samples is done by demonstrating the presence of type- and subtype-specific antigens (see below).

monoclonal antibody technology to demonstrate type A influenza antigens in a solid-phase, flow-through ELISA. This test will detect ATV antigens in AAF or directly in clinical specimens (cloacal and tracheal/orophaiyngeal swabs) from birds using a 15 min procedure (31). These test kits have potential use for rapid screening of poultry flocks for avian influenza (Al), but vary in sensitivity, false positive rate and the chickens must be in the acute stage of the disease, that is, showing clinical respiratory or systemic signs or be dead, to provide sufficient viral antigen (minimum of 1055 EIDso/ml of AAF) for detection. An experimental antigen-capture ELISA (ACELISA) using microtiter plates will also detect type A influenza viral RNP in cloacal and tracheal/oropharyngeal swabs (11). However, in waterfowl surveys, presumably because of low virus titers, AC-ELISA had low sensitivity and specificity as compared to virus isolation in chicken embryos (3). Visual Demonstration of AIV Antigen. Demonstration of ATV antigen in tissue by immunocytochemistry, immunohistochemistry, or immunofluorescent microscopy has value for rapid diagnosis of Al, especially HPAI, but should be utilized only as an adjunct to virus isolation. DFA staining of acetone-fixed tissue impression smears can give a presumptive diagnosis of Al in less than 1.5 hr (4). Impression smears of liver and kidney are easiest to prepare and give the most consistent results, especially for diagnosis of HPAI. However, impression smears of tracheal mucosa and cloacal bursa also provide satisfactory results and are necessary samples to diagnose LPAI. ATV can also be demonstrated in fresh-frozen sections by DFA, IFA, and IP tests (20) and in formalin-fixed tissue sections by immunohistochemistry (7,20). However, formalin fixation reduced the ability %of immunofluorescent and immunochemistry procedures to detect AIV. Detection of antigen in kidney tissue sections by the IP test was consistent only when virus titers were 105 ELD5()/g of tissue or higher. The IFA and IP tests are considered to be more sensitive than the DFA test. Molecular Detection of Type-specific RNA. For those laboratories routinely testing for Al by RRT-PCR, this assay can also be used to detect presence of ATV in AAF, vaccines or clinical specimens utilizing primers/probes specific for RNA that codes for the highly conserved M protein, a type-specific protein, or H5, H7 or other HA subtypes if available.

Type Classification Influenza viruses are grouped into three types (A, B, or C) based on highly conserved internal components of the virion, namely the M and RNP proteins or genes. All avian, swine, and equine influenza viruses share common type A M and RNP proteins/genes that can be identified by any of four methods: 1) the agar-gel immunodiffusion (AGID) test with processed AAF or chorioallantoic membranes (CAMs) which is the most commonly used to identify AIV; 2) an antigen-capture solid-phase or flowthrough enzyme-linked immunosorbent assay (ELISA) with AAF or suitable clinical specimens; 3) visualization by the direct fluorescent antibody (DFA) and indirect fluorescent antibody (IFA) or immunoperoxidase (IP) tests with tissues or CAM smears and frozen sections; and 4) detection of the M gene of ATV in clinical specimens, AAF or vaccines by RT-PCR or assays. The AGID Test to Detect AIV Antigen. The procedure utilizes influenza A-positive and negative control sera, influenza A antigen, and an unknown sample (antigen). Either AAF or CAM from embryonating eggs can be the unknown antigen, but CAM is preferred because of higher virus content. Briefly, CAMs should be rinsed in phosphate-buffered saline (PBS) (pH 7.2), pooled, and carefully drained, opening pockets to remove excess fluid, before they are ground in an electric blender or hand-operated tissue grinder and used (undiluted) as antigen in the AGID test (6). Alternatively, undiluted AAF can be used as the unknown antigen, but the infectivity titer probably needs to be at least ΙΟ5-106 mean embryo lethal dose (ELD50) to give satisfactory results (15). If the undiluted AAF is negative for influenza A antigen by the AGID test, it may be necessary to concentrate the AAF by acid precipitation of the antigen to eliminate the possibility of a false negative result because of a low virus titer. Briefly, eight to 10 ml of AAF is acid precipitated (IN HC1, pH 4.0, 4 C) for 60 min in an ice bath. The precipitate is centrifuged at 1000 x g for 10 min at 4 C, the supernatant is discarded, and the pellet is resuspended in 80100 μΐ glycine-Sarkosyl buffer. The resulting antigen is tested by the AGID test. The AGID test can be performed in Petri dishes with 0.9% agarose (ME grade) in PBS (0.01 M, pH 7.2) with an additional 8% NaCl. A seven-well template with a center well surrounded by six evenly spaced wells is used. The wells are 2.4 mm apart and 5.3 mm in diameter. The precipitin line will be present within 24 hr at room temperature and may increase in density for up to 48 hr. If a precipitin line develops between the suspect antigen and the positive serum, and if that line is continuous with the line between the adjacent positive antigen and the antiserum, the agent can be identified as type A influenza virus. Antigen Capture to Detect AIV Antigen. Detection of ATV in samples can be done by demonstrating the presence of specific influenza viral antigens with colorimetric assays. Commercially licensed test kits for human and veterinary medical use such as Directigen Flu A® (Becton Dickinson Microbiology Systems, Sparks, MD), NOW Flu A® (Binax, Portland, ME) Flu Detect® (Synbiotics, San Diego, CA) and others are available that use

Subtype Classification Once an isolate or viral proteins have been identified as type A influenza by either the AGID test, solid-phase ELISA, AC-ELISA, or RT-PCR, the next step is H and N subtyping. There are currently 16 Η (H1-H16) and 9 N (N1-N9) subtypes recognized as determined by the HI (15) and neuraminidase-inhibition (NI) (32) tests, respectively. The HI and NI tests are performed in microtiter plates with monospecific reference sera directed against whole virus preparations or purified H and N antigens. Alternatively, molecular subtyping assays, utilizing, have been developed for North American lineages of H5 and H7 ATV (22) but are not yet available for the other subtypes. All isolates identified as subtypes H5 and H7 are Notifiable Avian Influenza (NAT) and should be reported to state and federal authorities. The H5 and H7 NAI can be LP or HPAI viruses. Subtype determination is beyond the scope of most diagnostic laboratories not specializing in influenza viruses. Assistance is available from the National Veterinary Services Laboratories (NVSL), Ames, Iowa. The NVSL is a reference laboratory of the World Organization for Animal Health (OIE). The HI Test to Subtype AIV Isolates. Four HA units of the newly isolated virus can be tested against sera of known H subtypes with a suspension of 0.5% chicken erythrocytes (see Chapter 47). However, because of antigenic differences between ATV subtypes and slight differences even within subtypes, the AIV should be identified only by experienced personnel using monospecific antisera in reference laboratories. 131

David E. Swayne, Dennis A. Senne and David L. Suarez

With HI tests for ATV, care must be taken in selecting reagents used to identify the H subtype of the virus to avoid problems with

steric hindrance. Steric hindrance can occur if the antiserum used for H subtyping also contains N antibodies that are homologous

Table 29.1. Amino acid sequences at the HAO cleavage site of H5 and H7 AIV strains (modified from Senne et al., 1996 (18). Virus Strain

Subtype

Pathotype

HAO Cleavage site sequence

A/Chicken/Hidalgo/26654-1368/94

H5N2

LPAI

A/Emu/NY/12716-67/94

H5N9

LPAI

PQ------------ ....................RETR GLF * PQ------------ --------------- RETR GLF *

A/Turkey/CA/6878/79

H5N3

LPAI

PQ------------ ................... RETR GLF *

A/Chicken/Queretaro/14588-19/95

H5N2

HPAI

PQ------------ -------- RKRKTR GLF *

A/Γurkey/England/91

H5N1

HPAI

PQ------------ --------- RKRKTR GLF *

A/Γ urkey/Ireland/83

H5N8

HPAI

PQ------------ --------- RKRKKR GLF *

A/Chicken/Hong Kong/220/97

H5N1

HPAI

PQ------------ —RERRRKKR GLF *

A/Chicken/NY/13142-5/94

H7N2

LPAI

p E------------ ------------ NPKTR GLF *

A/Turkey/Italy/977/99

H7N1

LPAI

P £------------ ------------- 1 PKG R G *

LF

A/Quail/AR/16309-7/94

H7N3

LPAI

PE------------ ----------- -N PKT R G *

LF

A/Chicken/Chile/176822/02

H7N3

LPAI

PE

------------ K PKT R G *

LF

A/Chicken/Canada/AVF VI/04

H7N3

LPAI

GLF * PE.................------------ NPKTR

A/Chicken/Victoria/85

H7N7

HPAI

p e------------ ---- IPKKREKR GLF *

A/Chicken/The Netherlands/04

H7N7

HPAI

GLF * PE------------ ------- 1 PKRRRR

A/Turkey/Italy/4580/99

H7N1

HPAI

G * P E................. —IPKGSRVRR

LF

A/Chicken/Chile/4325/02

H7N3

HPAI

PEKPKTCSPLSRCRKT R G *

LF

A/Chicken/Canada/AVFV2/04

H7N3

HPAI

PE----- NPKQA YQKRMTR*GLF

* indicates cleavage site, dashes are used for alignment. Basic amino acids arginine (R) and lysine (K) are shown in bold. with the unknown isolate. The specific reaction of the N antibodies can interfere nonspecifically with the H, leading to nonspecific inhibition and possible misidentification of an isolate (17). The NI Test to Subtype AIV Isolates. The NI test is used to identify the other important surface antigen of the influenza viruses (8,32). This test, being more complex than the HI test, is generally performed only by those engaged actively in influenza research or influenza virus characterization. All nine of the known influenza N subtypes have been identified in avian species.

site of H protein. The virulence marker for HPAI is characterized by the presence of multiple basic amino acids at the H cleavage site (Table 29.1); however, caution should be exercised because some recent HPAI viruses have been shown not to have this virulence marker (10,24). The method by which the presence of multiple basic amino acids at the cleavage site can be determined has largely been limited to RT-PCR amplification of the coding sequences surrounding the cleavage site of the H protein, followed by sequencing of the amplicon by automated sequencing methods (18,33). This approach has been used to obtain sequences from a large number of H5 and H7 subtype viruses representing different temporal, geographical and host origins. However, as with any molecular based assay, success in the amplification of the target sequence will depend on the sensitivity of the primers used. Phylogenetic Analysis As part of the routine characterization of AIV from new outbreaks, the viruses will be at least partially sequenced for the hemagglutinin and other viral genes. The sequence information, as mentioned previously can help determine the pathogenic potential for H5 and H7 viruses, and additionally can be used for molecular epidemiology purposes. The sequence data can be compared to the influenza sequence database either through a blast search of the National Center of Biological Information or by phylogenetic analysis using a variety of available programs. The phylogenetic analysis principally provides a graphical representation of the relationships or closeness of viruses. The NCBI blast search is a quick and inexpensive way to determine the sequence similarity to all the sequence information available in GenBank. Further information on the origins of a virus isolate may be gained by phylogenetic analysis. The determination of the origin of a virus is useful to understand how the outbreak may have occurred. For example, the Virginia LP H7N2 in 2002 affected 197 flocks and cost the government in excess of 60 million dollars. Phylogenetic analysis clearly showed that this virus was not a recent introduction from wild birds, but was related to the viruses that were being isolated from the live bird marketing system in the Northeast USA (21). This information helped provide the basis for the decision to

Pathogenicity Determination Chicken Pathogenicity Test. Assessment of pathogenicity of a newly isolated ATV is critical for appropriate control strategies and international trade. Precautions should be taken to assure adequate containment of the virus before attempting bird inoculations. Pathogenicity testing of new isolates should be conducted according to guidelines published by the OIE and the U.S. Animal Health Association (13). Procedures currently recommended include intravenous inoculation of eight 4 to 8-wk-old chickens, and, if the isolate is of the H5 or H7 subtype, determination of the amino acid sequence at the cleavage site of the H. Isolates that are lethal for six, seven or eight (>75%) of eight experimentally inoculated chickens, or have an intravenous pathogenicity index (IVPI) >1.2 in 6-wk-old susceptible chickens are considered HPAI. In addition, H5 or H7 subtype viruses that do not meet the >75% mortality criteria but have an amino acid sequence at the H cleavage site that is compatible with HPAI viruses are also classified as HPAI (see Molecular Assessment of Pathogenicity below). Isolates that do not meet the criteria to be classified as HP should be designated simply as ATV or LPAIV with its corresponding H and N subtype designation. Avian influenza virus isolates from any H subtype can be pathogenic, but to date, only H5 and H7 subtypes have been HP. However, the vast majority of H5 and H7 subtypes are LPAI. Molecular Assessment of Pathogenicity. An understanding of the molecular basis for pathogenicity of AIV has made it possible to predict virulence of some strains of H5 and H7 subtype viruses based on presence or absence of a virulence marker at the cleavage 132

Chapter 29 Avian Influenza

are species specific, although the available commercial kits will detect serologic response in both chickens and turkeys. Development of a competitive ELISA would result in a single test usable for all avian species. The HI Test for Antibody Detection in the Host. The HI test allows avian influenza H subtypes to be differentiated on the basis of the antigenic character of the H and is an essential follow-up test for AGID-positive sera samples. This subtype specificity makes the HI test of limited value in initial screening of suspect birds or flocks, unless secondary spread of a previously identified ATV is being monitored. A large battery of control sera and test antigens representing all 16 of the known H subtypes would be necessaiy for detecting primary infection of an unknown H subtype. However, that still would not rule out the possibility that a serum might contain antibodies to a new H subtype not yet described, or because of antigenic differences caused by antigenic drift. HI testing for subtyping purposes is best suited for references laboratories, especially because sera may need to be treated with a receptor­ destroying enzyme to eliminate nonspecific HI reactions (15). The NI Test for Antibody Detection in the Host. The NI test allows ATV to be differentiated on the basis of the antigenic character of the N and is an essential follow-up test for AGIDpositive samples. NI testing is best suited for reference laboratories.

eradicate the virus and helps us understand that the virus was well adapted to poultry and could spread easily from flock to flock. The minimum sequence information necessary for a reasonable tree is at least 300 base pairs, but the more sequence information available the more discriminatory the phylogenetic trees As sequencing has become cheaper and more available, the expectation is more to sequence all eight gene segments to provide the most comprehensive view of the isolate possible.

SEROLOGIC DETECTION IN THE HOST

Routine surveillance in individual birds or flocks for serologic evidence of ATV infections is important for early detection and regulatory surveillance. Four major tests are used: AGID, ELISA, HI and NI. Historically, the virus-neutralization (VN) test has been performed as is done with other viruses, such as NDV. Specificity is similar to an HI test, but the VN test is infrequently used for diagnostic purposes in poultry medicine. The AGID Test for Antibody Detection in the Host. The AGID is the preferred serologic surveillance test in the U.S.A, because a single test detects serologic response in all bird species and against infection by all type A influenza viruses. Type A AGID test reagents (antigen and positive serum) are readily available from the NVSL and a commercial company (Charles River-SPAFAS, Norwich, CT). Alternatively, CAMs can be processed as described previously in the section on the AGID test to detect ATV antigen and used as the test antigen (6). Briefly, the CAM is ground, frozen and thawed three times, and centrifuged at a low speed (500-1500 x g). The supernatant is removed and inactivated with formalin (0.1% final concentration). The antigen can be used immediately or after incubating at 37 C for 36 hr to assure virus inactivation. The CAM from three eggs should yield about 1 ml of antigen. The antigen can be stored at 4 C for several weeks or frozen indefinitely at -10 C. An antigen consisting principally of M protein can also be prepared from a virus suspension in AAF (15). Care should be taken to balance the concentration of antigen to match the antiserum so that the precipitin line forms midway between the antigen and antiserum well. Only antigen and antiserum that produces a single, narrow, bold line that extends completely into adjacent test wells (containing negative serum) should be used. Also, a set of known strong and weak positive serums and negative serums should be used to assure adequate sensitivity and specificity of the reagents. A protocol for preparation of antigen and antiserum is available from the NVSL. The AGID influenza type A test for antibody detection in the host utilizes unknown test sera, influenza A test antigen, and influenza A positive control antiserum. The precipitin line between the ATV test antigen and positive antiserum will be present within 24 hr at room temperature and will increase in density for up to 48 hr; however, interpretation of most test results can be made at 24 hr. Sera from infected chickens or turkeys will yield positive AGID results within 5-6 days and up to 3 mo after the appearance of clinical signs. Because not all sera will be of the same potency, 20 or more individual sera should be tested from a flock. All AGIDpositive sera samples should be subtyped by HI and NI tests. An ELISA for Antibody Detection in the Host. Several indirect ELISA test kits to detect type A group-specific RNP and M antibodies are commercially available (Synbiotics, San Diego, CA; IDEXX, Westbrook, ME). ELISA procedures have also been developed for internal use by diagnostic laboratories (1). The indirect ELISA is a reliable test amenable to semi-automation and the rapid survey of large numbers of samples, but test results should be interpreted on a flock and not an individual bird basis. Strong positive reactions on ELISA correlate well with AGID. However, reactions in the suspect range should be retested by AGID or HI tests to confirm ATV infections. The threshold for ELISA-positive reactions should be set to avoid false positives. Indirect ELISA tests

DIFFERENTIATION FROM CLOSELY RELATED AGENTS

The influenza viruses produce a wide range of clinical signs and lesions, and diagnosis depends upon viral and serologic identification, especially in the initial flocks. Isolates must be distinguished from all other hemagglutinating agents, including some avian adenoviruses, Newcastle disease virus (NDV), and other avian paramyxoviruses by use of the AGID test with known positive monospecific influenza A antiserum, by the use of ATV specific antigen capture tests, or by RT-PCR assays specific for ATV or NDV. Diagnosis can be complicated by the presence of other microorganisms, such as mycoplasmas. Dual isolations of influenza virus and Newcastle disease virus are not unusual. Standard laboratory techniques using specific antisera to neutralize other viruses or antibiotics to control bacterial growth are helpful in such situations. ACKNOWLEDGMENT

The authors thank CW. Beard for contributions to prior editions. REFERENCES

1. Adair, B.M, D. Todd, E.R. McKillop, and MS. McNulty. Detection of influenza A type-specific antibodies in chicken and turkey sera by enzymelinked immunosorbent assay. Avian Pathol. 18:455-463. 1989. 2. Alexander, D.J. A review of avian influenza in different bird species. Vet.Microbiol. 74:3-13. 2000. 3. Alfonso, C.P., B.S. Cowen, and H. Vancampen. Influenza A viruses isolated from waterfowl in two wildlife management areas of Pennsylvania. J.Wildl.Dis. 31:179-185. 1995. 4. Allan, G.M and MS. McNulty. A direct immunofluorescence test for the rapid detection of avian influenza virus antigen in tissue impression smears. Avian Pathol. 14:449-460. 1985. 5. Bankowski, R.A. Introduction and objectives of the symposium. In: Proceedings of the First International Symposium on Avian Influenza, RA Bankowski, ed. U.S. Animal Health Association, Richmond, Virginia, pp. vii-xivl981. 6. Beard, C.W. Demonstration of type-specific influenza antibody in mammalian and avian sera by immunodiffusion Bull.World Health Organ. 42:779-785. 1970. 7. Brown, C.C., H.J. Olander, and D A. Senne. A pathogenesis study of highly pathogenic avian influenza virus H5N2 in chickens, using immunohistochemistry. J.Comp.Pathol. 107:341-348. 1992.

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8. Deusen, R.A., V.S. Hinshaw, D.A. Senne, D. Pellacani, and R.A. VanDeusen. Micro neuraminidase-inhibition assay for classification of influenza A virus neuraminidases. Avian Dis. 27:745-750. 1983. 9. Elbers, AR.W, T.H.F. Fabri, T.S. de Vries, J.J. de Wit, A Pijpers, and G. Koch. The highly pathogenic avian influenza A (H7N7) virus epidemic in The Netherlands in 2003--lessons learned from the first five outbreaks. Avian Dis. 48:691-705. 2004. 10. Hirst, M, C.R. Astell, M Griffith, S.M Coughlin, M Moksa, T. Zeng, D.E. Smailus, R.A. Holt, S. Jones, MA. Mana, M Petrie, M Krajden, D. Lawrence, A Mak, R. Chow, D.M Skowronski, Tweed S Aleina, S. Goh, R.C. Branham, J. Robinson, V. Bowes, K. Sojonky, S.K. Byme, Y. Li, D. Kobasa, T. Booth, and M Paetzel. Novel avian influenza H7N3 strain outbreak, British Columbia. Emerging.infectious. diseases. 10:2192-2195. 2004. 11. Kodihalli, S., V. Sivanandan, D.A. Halvorson, K.V. Nagaraja, and MC. Kumar. Antigen-capture ELISA for rapid diagnosis of avian influenza virus in commercial turkey flocks. Journal of Veterinary Laboratory Diagnosticians 5:438-440. 1993. 12. Lee, C.W., D.E. Swayne, J.A. Linares, D.A. Senne, and D.L. Suarez. H5N2 avian influenza outbreak in Texas in 2004: the first highly pathogenic strain in the United States in 20 years? J. Virol, in press2005. 13. OIE. International Animal Health Code. http://www.oie.int/eng/normes/MCodeZA 00003.htm 2002. 14. OIE. Highly pathogenic avian influenza in South Africa. OIE Disease Information 17(33):http://www.oie.int/eng/info/hebdo/AIS 32.HTM2005. 15. Palmer,D.F., M.T.Coleman, W.D.Dowdle, and G.O.Schild. Advanced laboratory techniques for influenza diagnosis. Immunology series no. 6. U.S. Department of Health, Education and Welfare, Public Health Service, Centers of Disease Control, Atlanta, Georgia. 1975 16. Rojas, H., R. Moreira, P. Avalos, I. Capua, and S. Marangon. Avian influenza in poultry in Chile. Vet.Rec. 151:1882002. 17. Schulman, J.L. and E.D. Kilbourne. Independent variation in nature of hemagglutinin and neuraminidase antigens of influenza virus: distinctiveness of hemagglutinin antigen of Hong Kong-68 virus. Proc.Natl.Acad.Sci.U.S.A. 63:326-333. 1969. 18. Senne, D.A., B. Panigrahy, Y. Kawaoka, J.E. Pearson, J. Suss, M Lipkind, H. Kida, and R.G. Webster. Survey of the hemagglutinin (HA) cleavage site sequence of H5 and H7 avian influenza viruses: amino acid sequence at the HA cleavage site as a marker of pathogenicity potential. Avian.Dis. 40:425-437. 1996. 19. Siemens, RD., R.S. Cooper, and J.S. Orsbom. Isolation of type-A influenza viruses from imported exotic birds. Avian Dis. 17:746-751. 1973.

20. Slemons, R.D. and D.E. Swayne. Replication of a waterfowl-origin influenza virus in the kidney and intestine of chickens. Avian Dis. 34:277284. 1990. 21. Spackman, E., D.A. Senne, S. Davison, and D.L. Suarez. Sequence analysis of recent H7 avian influenza viruses associated with three different outbreaks in commercial poultry in the United States. J Virol. 77:1339913402. 2003. 22. Spackman, E., D.A. Senne, T.J. Myers, L.L. Bulaga, L.P. Garber, ML. Perdue, K. Lohman, L.T. Daum, and D.L. Suarez. Development of a real­ time reverse transcriptase PCR assay for type A influenza virus and the avian H5 and H7 hemagglutinin subtypes. J.Clin.Microbiol. 40:3256-3260. 2002. 23. Suarez, D.L. and C.S. Schultz. Immunology of avian influenza virus: a review. Dev.Comp.Immunol. 24:269-283. 2000. 24. Suarez, D.L., D.A Senne, J. Banks, I.H. Brown, S.C. Essen, C.W. Lee, RJ. Manvell, C. Mathieu-Benson, V. Moreno, J.C. Pedersen, B. Panigrahy, H. Rojas, E. Spackman, and D.J. Alexander. Recombination resulting in virulence shift in avian influenza outbreak, Chile. Emerging Infectious Diseases 10:693-699. 2004. 25. Suarez, D.L., P.R. Woolcock, A.J. Bermudez, and D.A Senne. Isolation from turkey breeder hens of a reassortant H1N2 influenza virus with swine, human, and avian lineage genes. Avian Dis. 46:111-121. 2002. 26. Swayne, D.E. Microassay for measuring thermal inactivation of H5N1 high pathogenicity avian influenza virus in naturally-infected chicken meat. International Journal of Food Microbiology In press:2006. 27. Swayne, D.E. and J.R. Beck. Heat inactivation of avian influenza and Newcastle disease viruses in egg products. Avian Pathol. 33:512-518. 2004. 28. Swayne, D.E. and D.A. Halvorson. Influenza. In: Diseases of Poultry, 11th ed. Y.M Saif, H.J. Barnes, AM Fadly, J.R. Glisson, L.R. McDougald, and D.E. Swayne, eds. Iowa State University Press, Ames, LA. pp. 135-160. 2003. 29. Swayne, D.E. and RD. Slemons. Evaluation of the kidney as a potential site of avian influenza virus persistence in chickens. Avian Dis. 36:937-944. 1992. 30. Tang, Y., C.W. Lee, Y. Zhang, D.A Senne, R. Dearth, B. Byrum, D.R. Perez, D.L. Suarez, and Y.M Saif. Isolation and characterization of H3N2 influenza A virus from turkeys. Avian Dis. 49:207-213. 2005. 31. Vakharia, V.N., E.T. Mallinson, and P.K. Savage. A 15-min test for Al. Broiler Industry July:42-46. 1996. 32. Webster, R.G. and C.H. Campbell. An inhibition test for identifying the neuraminidase antigen on influenza viruses. Avian Dis. 16:1057-1066. 1972. 33. Wood, G.W., J.W. McCauley, J.B. Bashiraddin, and D.J. Alexander. Deduced amino acid sequences at the haemagglutinin cleavage site of avian influenza A viruses of H5 and H7 subtypes. Arch.Virol. 130:209-217. 1993.

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30 NEWCASTLE DISEASE VIRUS AND OTHER AVIAN PARAMYXOVIRUSES Dennis J. Alexander and Dennis A. Senne SUMMARY. Avian paramyxoviruses are members of the family Paramyxoviridae, genus Avulavirus. Nine serotypes are recognized and are termed APMV-1 (which is synonymous with Newcastle disease virus [NDV]) to APMV-9. APMV-1 viruses naturally infect a large range of avian species, but other avian paramyxoviruses appear to be restricted in their host distribution. APMV-1, APMV-2, APMV-3, APMV-6 and APMV-7 viruses have produced significant disease in naturally infected poultry. In poultry, NDVs can cause a range of clinical signs from subclinical to sudden very high mortality, depending on virus strain and host. Virulent NDV is a World Organization for Animal Health (OLE) listed notifiable disease and subject to international reporting. Agent Identification. Diagnosis of all avian paramyxoviruses is by isolation and identification of the virus. Virus isolation is usually done by the inoculation of embryonating chickens’ eggs. For NDVs, it is necessary to test for pathogenicity to determine if statutory control measures should be enforced. Vaccination of poultry and other birds for Newcastle disease is practiced throughout the world and vaccination for APMV-3 infections in turkeys is done in some countries. For NDVs, tests based on RT-PCR have been developed that facilitate detection of the virus in clinical specimens from infected animals; in addition a molecular-based assessment of virulence is also available. Serologic Detection in the Host. Serologic tests have limited diagnostic value in vaccinated poultry, but in unvaccinated birds, may provide a rapid early indication of infection. Caution should be exercised in relating positive serology to disease signs in view of the subclinical nature of infections of birds with most avian paramyxoviruses and because of cross-reactions that may occur between serotypes.

in chickens and turkeys may cause few signs of disease in ducks. The clinical signs that may be associated with ND are respiratory distress; diarrhea; cessation of egg production; depression; edema of head, face, and wattles; nervous signs; and death. Some, all, or none of these signs may be present. Avian paramyxovirus type 2 and APMV-3 infections of turkeys are common in some countries (including the United States) and are usually associated with respiratoiy disease and egg production problems. APMV-2 viruses have afso been reported in chickens. APMV-3 viruses have been associated with nervous disease with high mortality in some captive psittacine species. APMV-6 and AMPV-7 virus infections in turkeys have been associated with respiratory signs and elevated mortality. APMV-5 viruses have been isolated from budgerigars and lorikeets in association with invariably lethal disease. All other isolations of other serotypes of avian paramyxoviruses have been made from commercial or feral birds with no associated disease.

INTRODUCTION Three virus families, Rhabdoviridae, Filoviridae and Paramyxoviridae, form the order Mononegavirales. These are viruses with negative-sense, single-stranded, nonsegmented, RNA genomes. The family Paramyxoviridae has two subfamilies. The subfamily Pneumovirinae consists of two genera, Pneumovirus, which includes the mammalian respiratory syncytial viruses and Metapneumovirus which includes the avian pneumovirus responsible for turkey rhinotracheitis and swollen head syndrome. The subfamily Paramyxovirinae consists of five genera. The genus Morbillivirus includes measles, rinderpest, and the distemper viruses. The genus Paramyxovirus includes Sendai virus and other mammalian parainfluenza viruses. The genus Henipavirus which is formed from the recently discovered Nipah and Hendra viruses. The genus Rubulavirus includes mumps virus and human parainfluenza viruses 2 and 4. Newcastle disease (ND) virus (APMV-1), and the other avian paramyxoviruses (APMV-2 to APMV-9) are placed in the genus Avulavirus (16). The name avian paramyxoviruses has been retained rather than adopting the name avulaviruses. It should be noted that avian paramyxovirus type 1 (APMV-1) and ND virus (NDV) are synonymous, although the term pigeon paramyxovirus type 1 (PPMV-1) has been used to distinguish the antigenic variant APMV-1 virus responsible for the continuing panzootic in racing and other types of pigeons. Nine distinct serotypes of avian paramyxovirus have been defined using standard serologic tests (6). The prototype strains are listed in Table 30.1. Avian paramyxovirus type 1 (or ND) viruses are important pathogens for birds of all types, and in most countries infection with virulent forms is a notifiable disease, suspicion of which requires immediate notification of animal health authorities. It is an OIE listed notifiable disease. NDV has a worldwide distribution, although in some countries, only viruses of low virulence for chickens have been reported. NDV has a large host range and has been reported to infect more than 240 species of birds, probably all species of bird are susceptible to infection.

SAMPLE COLLECTION

For virus isolation For all avian paramyxoviruses, the samples associated most consistently with successful virus isolation have been feces or cloacal swabs. Viruses have also been isolated frequently from the respiratory tract using tracheal/orophaiyngeal swabs. Thus, it is important when taking samples for virus isolation that tracheal/oropharyngeal swabs and cloacal swabs or fecal material are included regardless of the clinical signs or the postmortem lesions. Additional samples collected from dead birds should reflect the clinical signs (e.g., brain if neurologic signs were evident) and the obviously affected organs. Virulent NDV is commonly isolated from lung, spleen, liver, heart, and brain. Ideally, all samples should be dealt with separately. In practice, it is usual to make pools of the organs and tissues, but to treat tracheal/oropharyngeal swabs and fecal samples separately, both from each other and from organ and tissue samples. However, tissues/organs from different birds should not be pooled, to avoid the possibility of neutralization of ND virus from one bird by specific antibodies from another bird. For birds showing neurologic signs before death, especially when investigating APMV-1 infections in pigeons, it is advisable to process brain samples separately from other organs and tissues. In NDV vaccinated flocks suspected of being infected with virulent NDV, sampling of daily mortalities has been shown to be more productive in detecting the virus than the random sampling of birds in the flock. Swabs should be placed in sufficient antibiotic medium to ensure full immersion. Fecal samples and finely minced tissues or organs

CLINICAL DISEASE

The clinical signs seen in birds infected with NDV vary widely and are dependent on factors such as the virus strain, host species, age of host, presence of other organisms, environmental stress, and the immune status of the host. In chickens the disease caused by different strains of NDV may vary from sudden death with 100% mortality to subclinical infection. The influence of host species may be equally marked and, for example, viruses causing severe disease

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should be placed in antibiotic medium as 10-20% w/v suspensions. The antibiotic medium should be based on phosphate-buffered saline at pH 7.0-7.4 (checked after the addition of antibiotics). Protein based media e.g. brain heart infusion (BHI) or tris-buffered tryptose broth (TBTB) have also been used and may give added stability to the virus, especially during shipping. The antibiotics used and their concentrations may be varied to suit local conditions and availability. Very high levels of antibiotics may be necessary for fecal samples; suggested levels are: 10,000 IU/ml penicillin, 10 mg/ml streptomycin, 0.25 mg/ml gentamycin, and 5,000 IU/ml nystatin. These levels can be reduced up to fivefold for tissues and tracheal swabs. If control of Chlamydophila is desired 0.05-0.1 mg/ml oxytetracycline should be included. Successful isolation is enhanced by rapid processing of samples. Virus infectivity in tissues or organs is destroyed by putrefaction and this should be avoided by refrigerating samples at 4 C or on ice during transport or storage. Placing samples in antibiotic medium for transportation should be done in addition to chilling and not as an alternative to chilling. Most avian paramyxoviruses appear to be able to survive in fecal samples for long periods even at relatively high ambient temperatures. Most NDVs will survive a single freeze-thaw cycle with little loss in infectivity, but this may not be the case with other avian paramyxoviruses.

PREFERRED CULTURE MEDIA AND SUBSTRATES

Virus isolation Newcastle disease virus and most other avian paramyxoviruses will grow well in a variety of cell culture systems. Avirulent NDVs and other avian paramyxoviruses usually require the addition of trypsin (0.005-0.01 mg/ml) to facilitate growth in cell cultures. However, the most sensitive method of isolation for all these viruses is the inoculation of embryonating chicken eggs. Samples in antibiotic media should be left at least 30 min at room temperature before inoculation (this step can be omitted if samples have been stored overnight or longer at -4 C). Suspensions of tissues and organs or swab washings should be centrifuged at 1000 x g for 10-20 min, and 0.1-0.3-ml volumes of the supernatant are inoculated into the allantoic cavity of four or five 9 to 11-day-old embryonating chicken eggs. Eggs should be obtained from a specific-pathogen-free (SPF) flock; if this is not possible, eggs obtained from a flock free of NDV antibodies may be considered. Use of eggs from antibody-positive flocks will reduce the ability of the virus to grow and the success of virus isolation. Inoculated eggs should be placed at 37 C and candled regularly. Eggs with dead or dying embryos and all eggs 5-7 days after inoculation should be chilled to 4 C, and the allantoic/amniotic fluids collected and tested for hemagglutinating activity. Ideally, negative fluids diluted 1:10 in antibiotic medium should be passaged at least once more. Some laboratories use undiluted allantoic fluid for the second and subsequent passages; no work has been done to show which method is the moje sensitive. To expedite diagnosis, some laboratories have used two 3 day passages and reported comparable results to two 6 day passages, but this has not yet been fully evaluated. Positive fluids need to be tested for freedom from bacteria. If bacteria are present the fluids may be passed through a 450-nm membrane filter or centrifuged to remove bacteria and repassaged in eggs after the addition of more antibiotics. Avian paramyxovirus type 5 viruses do not grow in the allantoic cavity of chick embryos, and several reports exist of greater success at isolating some other avian paramyxoviruses by inoculation into the yolk sac rather into than the allantoic cavity. For avian paramyxovirus isolations, consideration should be given to inoculation of material in to the yolk sac of 6 to 8-day-old chick embryos in addition to the allantoic cavity of 9 to 11-day-old eggs; this is essential for APMV-5 viruses.

For RT-PCR The use of real time RT-PCR (rRT-PCR) assays in diagnostic laboratories is becoming increasingly popular. A one step rRT-PCR assay has been shown to be highly sensitive in detecting NDV RNA in clinical samples (21) and has been used in the U.S. as a front line test in place of virus isolation in outbreaks of virulent ND and for surveillance. Samples suitable for the isolation of NDV, for the most part, also can be used for rRT-PCR (see collection of samples for virus isolation above). However, tracheal or oropharyngeal swabs are the specimens of choice because they contain the nucleic acid, are easy to process, and generally contain less extraneous organic material that can interfere with RNA recovery and amplification by PCR. Particular attention should be given to the selection of an appropriate method for RNA extraction to maximize recovery. Good quality RNA not containing PCR inhibiting substances is essential for successful amplification in the rRT-PCR assay. Also, use of an internal positive control (unrelated to the target cDNA) is highly recommended to monitor for presence of PCR inhibiting substances that could cause a false negative test result. Several commercial kits are suitable for the recovery of RNA from clinical specimens. The three commercial RNA extraction kits that have been most thoroughly evaluated for rRT-PCR assays are Trizol reagent (Invitrogen, Carlsbad, CA), RNeasy (Qiagen, Valencia, CA) and MagMAX (Ambion, Austin, TX). Trizol is an organic extraction method that works well for most specimens and has been shown to be best for extraction of RNA from suspensions of tissue. Although the Trizol kit will also work well for swab specimens, the toxic chemicals (phenol, guanidine isothiocyanate, and chloroform) required for this method makes it less desirable than other methods. The RNeasy extraction kit has been used extensively for processing tracheal and oropharyngeal swabs but has lower sensitivity for tissue suspensions and cloacal swabs, most likely because of the heavy organic load in these specimens; the extraneous RNA competes for binding sites in the extraction column, resulting in lower recovery rates of target RNA. The MagMAX kit can be performed in 96-well plates and therefore is more suitable than other extraction methods for high throughput testing. However, the suitability of this method for extraction of tissue suspensions and cloacal swabs has not yet been fully evaluated.

RT-PCR For NDV, most RT-PCR assays have been developed to aid in the identification or characterization of isolates by using primers that amplify portions of the genome related to a specific function, for example the fusion gene cleavage site and virulence (3, 10, 14, 19). Both conventional RT-PCR and nested RT-PCR assays have been used. More recently, a one step rRT-PCR assay has been developed to aid in the detection of NDV specific RNA in clinical samples as well as to provide a rapid method to distinguish between viruses of low and high virulence (21). This is especially important in outbreaks of virulent NDV where birds may be co-infected with both vaccine and virulent virus. In general, RT-PCR assays have four steps: isolation of the RNA; conversion of the viral RNA to cDNA by the use of reverse transcriptase; amplification of the cDNA by PCR; and evaluation of the PCR product (amplicon) for presence or absence of the target DNA sequence. Specificity of the amplicon is usually confirmed by electrophoresis of the amplicon in an agarose gel containing ethidium bromide, then comparing the size of the amplicon to markers of known molecular weight. However, other methods such, as enzyme restriction endonuclease assays and direct sequencing have been used to confirm specificity of the amplicon. The need for post-amplification processing of the amplican is one of the major

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drawbacks of using RT-PCR as a diagnostic assay because of the high potential for contamination of the laboratory and cross contamination of samples. Extreme precautions, such as the use of dedicated equipment and separation of sample processing activities from post amplification processing activities, must be taken to prevent cross contamination. Real time RT-PCR assays differ from conventional RT-PCR assays in that they require special thermocyclers and use fluorogenic hydrolysis (Taqman) probes or fluorescent dyes that, respectively, bind to specific targets on the PCR product or nonspecifically to any double stranded DNA molecule, to monitor for presence of target DNA after each PCR cycle, thus providing results in real time. The major advantages of the one step rRT-PCR assay is that results can usually be obtained in less than three hr and the elimination of the post amplification processing step, thus reducing concerns about cross contamination of samples. Unfortunately, many smaller laboratories have been hampered from using this technology because of the high start-up costs associated with purchasing the real time thermocyclers. The rRT-PCR assay described by Wise et al. (21) for detection of NDV RNA in clinical specimens was developed and validated during an outbreak of virulent ND in the U.S. in 2002-03 and eventually replaced virus isolation as the front line test during the outbreak. The rRT-PCR assay showed a sensitivity of 95% when results were compared to virus isolation on more than 1,400 specimens. The assay has three sets of primers and probes that are used in separate reactions: a matrix primer/probe set that is designed to detect most strains of NDV, a fusion primer/probe set that can identify virulent strains of NDV (including many PPMV-1 viruses) and a primer/probe set designed to detect low virulent strains of the virus. Samples are first screened with the matrix primers/probe then positive specimens are tested with the low virulent and fusion and primers/probe sets to confirm presence of low or highly virulent virus, respectively. At the peak of the outbreak, between 1,000 and 1,500 samples were tested daily by rRT-PCR. Testing capacity is mostly controlled by the extraction method used and availability of sufficient number of real time thermocyclers. In the U.S. the rRT-PCR assay has been authorized for use in more than 35 laboratories as a front line surveillance tool for NDV.

Newcastle Disease Virus and other Avian Paramyxoviruses

enzyme-linked immunosorbent assay (ELISA), for identifying and serotyping isolated viruses. However, the conventional method usually employed is the HI test, which is described in Chapter 47 on serological procedures. Serotype identification for avian paramyxoviruses is usually straightforward, although some levels of cross- reaction may be seen between the various serogroups. The most important of these is the cross-reaction seen in HI tests between APMV-1 and APMV-3 viruses, particularly when the latter are isolated from psittacines or other caged or exotic birds. Usually, confusion due to this relationship can be avoided if adequate control sera and antigens are used. Several laboratories have produced monoclonal antibodies (mAbs) to representatives of some of the avian paramyxovirus serotypes and these may be employed in HI tests or other tests to give highly specific serotyping of avian paramyxoviruses and even divisions within serotypes (see below). mAbs can be added to the initial battery of antisera used for virus identification to give maximum information about an isolate at an early stage. Morphology and Physicochemical Properties Avian paramyxoviruses are RNA viruses with helical capsid symmetry and have an nonsegmented, single-stranded genome of negative sense. The RNA has a molecular weight of about 5 x 106, which makes up about 5% by weight of the virus particle. Nucleotide sequencing of the complete NDV genome has shown it to consist of 15,156 nucleotides, although there may slight variations with different strains. The capsid of avian paramyxoviruses is assembled in the cytoplasm and becomes enveloped by modified cell lipoprotein membrane as the virus is budded from the cell surface. Two functional virus glycoproteins are inserted in the envelope, one (HN) possesses hemagglutination and neuraminidase activities, the other (F) is the fusion protein. During the infection process the HN protein is responsible for attaching the virus to the cell and the F protein brings about fusion between the cell and virus membranes to allow the genetic material to enter the cell. As seen by negative contrast electron microscopy, avian paramyxoviruses consist of pleomorphic particles that are usually rounded and 100-500 nm in diameter or are present as filamentous forms about 100 nm across. Surface projections on the envelope, approximately 8 nm long, represent the HN molecule, with the F molecules forming smaller projections. Virus envelopes are frequently disrupted so that a characteristic herring-bone nucleocapsid about 18 nm across may be seen emerging or free in virtually all paramyxovirus electron microscope preparations. The presence of nucleocapsid may serve as a useful method for distinguishing paramyxoviruses from influenza viruses, as the nucleocapsid of the latter is rarely ever seen.

AGENT IDENTIFICATION

Virus Identification Hemagglutination activity in bacteria-free fluids will be due to one of the nine avian paramyxovirus serotypes or one of the 16 influenza A virus subtypes. Most diagnostic laboratories will be primarily concerned with the presence of APMV-1 (NDV), which can be confirmed or ruled out by hemagglutination-inhibition (HI) tests with specific antiserum. Alternatively, procedures based on RT-PCR [see below] can be used to confirm the presence of APMV-1 (NDV) at this stage. In laboratories where differentiation of the avian paramyxovirus serotypes is done routinely, subjecting the hemagglutinating agent to HI tests against a range of specific antisera representing all the serotypes is usually considered most practicale. Antisera used for virus identification is generally prepared in chickens. Most laboratories have their own methods for the production of such antisera. For APMV-1 viruses of low virulence and other avian paramyxoviruses, suitable serum can usually be obtained by the infection of 6- to 9-wk-old SPF chickens with 0.1 ml of infectious allantoic fluid by injection into the leg muscle and nasal instillation of a further 0.1 ml of infectious allantoic fluid at the same time. This is repeated after 2 wk and the serum collected after another 3 wk. Specific polyclonal chicken sera can be used in most serologic tests, such as virus neutralization, agar gel immunodiffusion, and

Pathogenicity of NDV Isolates of NDV require further diagnostic characterization in addition to identification as APMV-1 serotype. Not only may different NDV strains show extremes of pathogenicity, but also viruses of low virulence are enzootic in feral birds in most countries. Use of live vaccines is almost universal, and exacerbation of infections with viruses of low virulence may mimic disease produced by highly virulent virus. Therefore, characterization to assess the virulence of the isolated virus is extremely important to ensure that the virus conforms to the definitions laid down in control and/or eradication policies. Pathotypes. Newcastle disease virus strains and isolates have been grouped into five clinico-pathological groups that relate to the disease signs and lesions produced in infected fully susceptible chickens (9): 1) viscerotropic velogenic NDV, which produces acute lethal infections in which hemorrhagic lesions are prominent in the gut; 2) neurotrophic velogenic NDV, which produces high

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mortality preceded by respiratory and neurologic signs (gut lesions are absent); 3) mesogenic NDV, which produces low mortality, acute respiratory disease, and nervous signs in some birds; 4) lentogenic NDV, which produces mild or inapparent respiratory infections; and 5) asymptomatic enteric NDV, which are avirulent viruses that appear to replicate primarily in the gut. These groups are not completely clear-cut and some overlapping between the signs associated with the different groups has been reported. If an NDV isolate is suspected to be virulent for chickens or other poultry species, it should be submitted to a reference laboratoiy for assessment. In the United States such viruses may be propagated only in a biosafety level 3 (BSL-3) laboratoiy, and similar or stricter restrictions apply in many other countries.

Thus there appears to be the requirement of a basic amino acid at residue 113, a pair of basic amino acids at 115 and 116 plus a phenylalanine at residue 117 if the virus is to be virulent for chickens (Table 30.3). The presence of these basic amino acids at these positions means that cleavage can be affected by a protease or proteases present in a wide range of host tissues and organs, but for lentogenic viruses, cleavage can occur only with proteases recognizing a single arginine, i.e. trypsin-like enzymes. Lentogenic viruses are therefore restricted in the sites where they are able to replicate to areas with trypsin-like enzymes, such as the respiratory and intestinal tracts, whereas virulent viruses can replicate and cause damage in a range of tissues and organs resulting in a fatal systemic infection. This molecular basis of virulence has been incorporated into the internationally recognized definition of viruses that cause ND for which control measures should be imposed and trade restrictions may be applied, as an alternative to the ICPI test. Defining such viruses as those where “Multiple basic amino acids have been demonstrated in the virus (either directly or by deduction) at the Cterminus of the F2 protein and phenylalanine at residue 117, which is the N-terminus of the Fl protein. The term ‘multiple basic amino acids’ refers to at least three arginine or lysine residues between residues 113 and 116. Failure to demonstrate the characteristic pattern of amino acid residues as described above would require characterization of the isolated virus by an ICPI test. [In this definition, amino acid residues are numbered from the N-terminus of the amino acid sequence deduced from .the nucleotide sequence of the F0 gene, 113-116 corresponds to residues -4 to -1 from the cleavage site.]” It is worth noting that all the so-called mesogenic viruses fall within this definition, possessing basic amino acids at the F0 cleavage site and having ICPI values >0.7. The method by which the presence of multiple basic amino acids at the cleavage site can be determined has largely been limited to RT-PCR amplification of the coding sequences surrounding the fusion protein cleavage site, followed by sequencing of the amplicon by automated sequencing methods (19). This approach has been used to obtain sequences from a large number of isolates of different temporal, geographical and host origins. However, as with any molecular based assay, success in the amplification of the target sequence will depend on the sensitivity of the primers used. Although the primers published by Seal et al. (19) work well for most NDV strains, they have shown limited sensitivity in amplifying viruses of genotype 6 isolated from ducks. Other methods for differentiating low from highly virulent NDV by RTPCR have been described (3, 10); however, they do not provide direct confirmation of the presence or absence of multiple basic amino acids at the fusion protein cleavage site

Pathogenicity Tests for NDV. Several pathogenicity tests have been developed to differentiate between NDV isolates of high and low virulence with some level of standardization. However, international agencies such as the World Organization for Animal Health [OIE] have adopted the intracerebral pathogenicity index [ICPI] test as the in vivo test used for defining the virulence of NDV isolates. Intracerebral Pathogenicity Index (ICPI). The ICPI is determined by inoculating 0.05 ml of a 1:10 dilution of infectious, bacteria-ffee allantoic fluid in sterile isotonic saline (antibiotics must not be present) into the brains of each of 10 1-day-old (24 to 40 hr-old) chicks from SPF parents. The birds are observed daily for 8 days and, at each observation, scored 0 if normal, 1 if sick, and 2 if dead. Birds that are sufficiently sick to be unable to eat or drink must be killed humanely and scored as dead at the next observation. The ICPI value is the mean score per bird per observation. Viruses very virulent for chickens give values approaching 2, lentogenic viruses give values close to 0 (Table 30.2). Some evidence exists that NDV isolates from unusual species may not show their true virulence for chickens in pathogenicity index tests until passaged several times in chickens. By internationally accepted definition, viruses that cause ND for which control measures should be imposed and trade restrictions may be applied are those with an ICPI of 0.7 or greater. When tested, other avian paramyxoviruses have usually produced indices similar to those of lentogenic NDV isolates. However, several reports have been made of psittacine APMV-3 viruses producing ICPI values of 1.0 or more, which emphasizes the need to differentiate APMV-1 viruses from other avian paramyxoviruses correctly.

Molecular Assessment of Pathogenicity. An understanding of the molecular basis that controls the virulence of NDV strains (18) has meant that it is now possible, using nucleotide sequencing techniques, to assess whether or not an isolate has the genetic make up to be highly pathogenic for poultry. The viral F protein brings about fusion between the virus membrane and the cell membrane so that the virus genome enters the cell and replication can begin. The F protein is therefore essential for replication, but during replication, NDV particles are produced with a precursor glycoprotein, F0, that has to be cleaved to Fl and F2 polypeptides, which remain bound by disulphide bonds, for the virus particles to be infectious. This post translation cleavage is mediated by host cell proteases. The cleavability of the F0 molecule has been shown to be related directly to the virulence of viruses in vivo. A large number of studies have confirmed the presence of multiple basic amino acids at the F0 cleavage site in virulent viruses. Usually the sequence has been 113RQK/RR F * 117 in virulent viruses, but most have had a basic amino acid at position 112 as well. In contrast, viruses of low virulence usually have the sequence 113K/RQG/ER * L 117.

Newcastle Disease Virus Strain Differentiation Using Monoclonal Antibodies Using conventional serologic techniques, NDV strains and isolates had been considered to form an antigenically homogeneous group. This meant that diagnosis usually resulted in little information relating to the source or the spread of the virus involved. MAbs may detect slight variations in antigenicity, such as single amino acid changes at the epitope to which the antibody is directed. Several groups of workers have developed mAbs against NDV strains, primarily for diagnostic or epidemilogical purposes. Some workers have used mAbs to distinguish between specific viruses, for example, two groups have described mAbs which distinguish between the common vaccine strains Hitchner Bl and La Sota (11, 17) whereas other mAbs have been used in distinguish vaccine

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Table 30.1. Avian Paramyxoviruses.

Prototype virus strain

Usual natural hosts

APMV-1/Newcastle disease virus

Numerous

APMV-2/chicken/Califomia/Yucaipa/56

Turkeys, passerines

APMV-3a/ turkey/Wisconsin/68 APMV-3A/parakeet/Netherlands/449/75

Turkeys Psittacines, passerines

APMV-4/duck/Hong Kong/D3/75 APMV-5Zbudgerigar/Japan/Kunitachi/74

Ducks Budgerigars, lorikeets

APMV-6/duck/Hong Kong/199/77

Ducks

APMV-7/dove/Tennessee/4/75 APMV-8/goose/Delaware/l 053/76

Pigeons, doves Ducks and geese Ducks

APMV-9/duck/New York/22/78

Disease produced in naturally infected poultry Very common, worldwide, varies from very severe to inapparent disease, depending on strain and host infected Common, probably worldwide, mild respiratory disease or egg production problems; severe if exacerbation occurs Turkeys only, in North America and Europe, mild respiratory disease but severe egg production problems worsened by exacerbating organisms or environment No infection of poultry reported None known No infection of poultry reported Mild respiratory disease and slightly elevated mortality in turkeys; none in ducks or geese Natural infections of turkeys with respiratory disease and ostriches have been reported No infection of poultry reported

None known

viruses from an epizootic virus in a given area (20). MAb typing was also used to establish the uniqueness of the variant NDV responsible for the pigeon panzootic and specific mAbs have proven particularly useful in identify the spread of this virus around the world. However, although use of mAb panels showed that viruses grouped on their ability to react with the same mAbs shared biological and epizootiological properties and that viruses tend to remain fairly well conserved antigenically during outbreaks or epizootics (7) this approach to epizootiological assessments of ND outbreaks has been largely supplanted by phylogenetic analysis.

SEROLOGIC DETECTION IN THE HOST Detection of an immune response to NDV or other avian paramyxoviruses may serve a useful diagnostic function, allowing detection of infections in unvaccinated birds and the monitoring of response to vaccination. Numerous serologic tests may be used to detect antibodies, but the most commonly used for this group of viruses is the HI test. Details of the procedures for this test are given in Chapter 46 on titration of biological suspensions.

Hemagglutination-Inhibition Suitably diluted infectious allantoic fluid may be used as antigen in HI tests for all avian paramyxoviruses except APMV-5. Use of noninfectious antigens usually is desirable; and for NDV formalininactivated virus (infectious allantoic fluid treated with 0.1% formalin) is often used as antigen. However, the hemagglutinin­ neuraminidase (HN) protein of some avian paramyxoviruses is not stable when treated with formalin and for those viruses betapropriolactone (0.05% final concentration) may be used as an alternative to formalin for inactivation. Hemagglutinating activity was not detected in the original APMV-5 isolates, but later viruses placed in this serotype have been shown to agglutinate red blood cells if concentrated following growth in chick embryo cell cultures. Some evidence exists that minor antigenic differences between strains of NDV can result in different HI titers with the same antiserum. This is particularly true using the variant PPMV-1 virus as antigen.

Phylogenetic analysis Nucleotide sequencing, after RT-PCR and phylogenetic analysis, has been used by a number of authors to assess genetic differences and genotypes of ND viruses. It has been established that sequences of as little as 250 base pairs give meaningful phylogenetic analyses, comparable with those obtained with much longer sequences (15, 19), which means that sequencing and phylogenetic analyses can be done rapidly. In the preliminary characterization studies of NDV, six lineages (I to VI) were determined using restriction enzyme analysis (8). These groupings have subsequently been confirmed, and two further lineages (VII and VHT) and several sublineages within these have been identified through nucleotide sequencing studies (13). Aldous et al., (4) studied the nucleotide sequences of a 375-nucleotide fragment at the 3’ end of the fusion protein gene, which includes the region encoding the nuclear localization signal sequence and the precursor fusion protein cleavage activation site, of 338 isolates of NDV representing a range of viruses of different temporal, geographical and host origins. They concluded that the isolates divided into six broadly distinct groups (lineages 1 to 6). Lineages 3 and 4 were further subdivided into four sublineages (a to d) and lineage 5 into five lineages (a to e). Essentially lineages 1, 2, 4 and 5 corresponded to the earlier defined lineages I, Π, VI and VH, with comparable sub-lineages but that the geno-groupings ΙΠ, IV, V, VUI corresponded to their sublineages (3a to 3d). In addition lineage 6 represent a new geno-group. Aldous et al., (4) proposed that genotyping of NDV isolates should become part of diagnostic virus characterization for reference laboratories by producing the 375-nucleotide sequence of the F gene routinely for all viruses and comparing the sequences obtained with other recent isolates and 18 viruses representative of the recognized lineages and sub-lineages. Such preliminary analysis should allow rapid epidemiological assessment of the origins and spread of the viruses responsible for ND outbreaks.

Interpretation of HI Response Serum titers will vary with the amount of antigen used in the test; figures quoted in this section assume 4 HA units of virus. For all avian paramyxovirus serotypes, birds that have not been immunized or infected usually have HI titers of less than 1:8, and nonspecific titers above this level are rare for most avian species. However, sera from other species may cause agglutination of chicken red blood cells to titers high enough to mask low positive inhibition. This agglutination, caused by the presence of natural serum agglutinins, can be removed by treatment of the sera with concentrated chicken red blood cells before testing. Alternatively, use of red blood cells from the homologous species can be considered. In the absence of vaccination, a positive specific HI response is confirmation of infection of the birds with that serotype. However, the possibility of cross-reaction between some avian paramyxoviruses should be considered, especially between viruses of APMV-1 (NDV) and APMV-3 serotypes. 139

Dennis J. Alexander and Dennis A. Senne

available commercially. The main advantage of ELISAs over more conventional tests, such as HI tests, is that they can be semi­ automated, enabling results to be obtained rapidly and inexpensively, especially when sera are to be screened for antibodies to several viruses (see Chapter 47 on serologic procedures). As a result, ELISAs have often become the method of choice for flock screening programs, particularly those aimed at assessing vaccine response. Studies aimed at standardizing ELISA tests and comparing them with conventional tests, for example Adair et al., (1), have concluded that ELISAs for NDV antibodies usually show good reproducibiEty, high comparative sensitivity and specificity, and correlate well with the HI test. The ELISAs should be modified and validated for species on which it is being used. ELISA tests are usually extremely sensitive and this may restrict their value for diagnostic testing when antigenic relationships exist between different viruses, which is the case with avian paramyxoviruses. Both these problems could be addressed by use of competitive or blocking ELISAs employing one or more mAb to NDV, although mAb(s) should be evaluated to determine that they react with all strains of NDV and not other avian paramyxoviruses.

Inactivated vaccines for APMV-3 viruses are used in the United States and Europe and use of APMV-2 vaccines has been undertaken in some countries. Little is known of the interpretation of the immune response to vaccines for these serotypes. Although considerable work has been done in attempts to assess the likely outcome of infection with virulent NDV at different levels of immunity, care must be taken in making such assessments due to the effects other factors may have. In broad terms titers at the low end of those measurably positive may protect against death in Table 30.2. Pathotypes and intracerebral pathogenicity indices__________ Range of indicesA

Pathotype

Examples of viruses®

ICPI

Viscerotropic velogenic Neurotropic velogenic

1.5-2.0

Herts 33, N.Y. Parrot 70181, CA2089/72

1.5-2.0

Texas GB

Mesogenic

1.0-1.5

Lentogenic

0.2-0.5

Asymptomatic enteric

0.0-0.2

*,Roakin *, Komarov *, Mukteswar * H Hitchner Bl , * La *, Sota Clone 30 * Ulster 2C *, *, V4 MC110

DIFFERENTIATION FROM CLOSELY RELATED AGENTS

A ICPI = intracerebral pathogenicity index in 1-day-old chicks, See text for details. B Strains marked with an asterisk are used routinely as vaccines in some countries.

Because of their abiEty to cause agglutination of red blood cells, usually without concentration or treatment, problems in differentiating avian paramyxoviruses are primarily from each other and from influenza viruses. Preliminary distinction from influenza viruses can be done by examination in the electron microscope where negative contrast staining will reveal morphologic differences, most notably the presence of the characteristic nucleocapsid emerging from disrupted paramyxovirus particles. The abiEty of paramyxoviruses, but not influenza viruses, to cause hemolysis of chicken red blood ceUs at pH 7.0 has also been used in the past. Influenza A viruses can be confirmed in allantoic fluid by the agar gel immunodiffusion test with type A positive antiserum, or by the use of antigen capture immunoassays (membrane bound or flow-through) designed to detect influenza A viruses. The antigen capture immunoassay tests are not Ecensed for veterinary use but have been shown to detect all 16 subtypes of avian influenza virus (ATV) and are highly specific. Differentiation between APMV-1 and ATV can also be done by the use of RT-PCR (conventional and real time), with primers directed to the highly conserved matrix protein genes of both APMV-1 and ATV. RT-PCR assays have also been developed that can discriminate between virulent and vaccine virus. However, it is recommended for definitive identification that isolates be submitted to a reference laboratory that can that can use a battery of polyclonal sera against each of the paramyxoviruses, or initially against the most likely serotypes, in HI tests. Differentiation of avian paramyxoviruses into serotypes can be more of a problem due to antigenic relationships between viruses of different serotypes, which are apparent in most conventional tests. This cross reaction in serologic tests is most marked between APMV-1 and APMV-3 viruses (particularly APMV-3 viruses isolated from psittacines). Usually the use of adequate control antigens and antisera leaves tittle doubt in serotyping isolates from these groups, but mAbs specific for APMV-1 and APMV-3 have been employed to simplify differentiation (5). The development and application of RT-PCR or rRT-PCR assays has made it possible to differentiate quickly NDV from other avian paramyxoviruses based on genetic differences and to detect mixtures of low and highly virulent ND virus in the same sample. However, such assays are not yet available to confirm the presence or distinguish between the other avian paramyxoviruses.

uncomplicated infections. However, even high titers will not prevent some replication of challenge virus and significant egg production losses may occur in flocks with prechallenge mean titers as high as 1:256. With most lentogenic live NDV vaccines, HI titers of 1:16 to 1:64 are usually obtained following a single dose. Repeated vaccination with similar or more virulent Eve virus vaccines may increase the immune response considerably, and if oil emulsion inactivated vaccines are included, titers up to 1:1024 to 1:4096 are common. Multiple vaccinations with either Eve-, killedor combinations of both virus vaccines wiU also increase the level of crossreactivity between serotypes of APMVs in the HI test. Enzyme-linked Immunosorbent Assays for NDV Antibodies Numerous ELISA tests have been developed for the detection of antibodies to NDV and several ELISA kits for this purpose are Table 30.3. Amino acid sequences at the F0 cleavage site of APMV-1 viruses modified from Aldous & Alexander (2)

Herts 33

velogenic

F0 Cleavage site amino acids 111 to 119 FIG* -GRRQRR

Essex ‘70

velogenic

FVG* -GRRQKR

Texas GB

velogenic

FIG* -GRRQKR

617/83

PPMV-1

FIG* -GGRQKR

34/90

velogenic

FVG* -GKRQKR

Beaudette C

mesogenic

FIG* -GRRQKR

Roakin

mesogenic

FIG* -GRRQKR

La Sota

lentogenic

LIG* -GGRQGR

Virus strain

Pathotype

Hitchner Bl

lentogenic LIG* -GGRQGR asymptomatic D26 LIG* -GGKQGR enteric asymptomatic MC110 LIG* -GERQER enteric asymptomatic 1154/98 LIG* -GRRQGR enteric note that all virulent viruses have phenyalalanie (F) at position 117, the Fl N-terminus

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11. Erdei, J., K. Bachir, E. F. Kaleta, K. F. Shortridge, and B. LommoL Newcastle disease vaccine (La Sota) strain specific monoclonal antibody. Arch. Virol. 96:265-269. 1987. 12. Hanson, R. P., and C. A. Brandly. Identification of vaccine strains of Newcastle disease virus. Science 122:156-157. 1955. 13. Herczeg, J., S. Pascucci, P. Massi, M Luini, L. Selli, I. Capua, I. and Bl Lomniczi. A longitudinal study of velogenic Newcastle disease vnws genotypes isolated in Italy between 1960 and 2000. Avian Pathol. 30: 163168. 2001. 14. Jestin, V., and A. Jestin. Detection of Newcastle disease virus RNA in infected allantoic fluids by in vitro enzymatic amplification (PCR). Arch. Virol. 118:151-161. 1991. 15. Lomniczi, B., E. Wehmann, J Herczeg, A. Ballagi Pordany, E F. Kaleta, O. Werner, G. Meulemans, P. H. Jorgensen, A. P. Mante, A. L Gielkens, I. Capua, and J. Damoser. Newcastle disease outbreaks in recent years in western Europe were caused by an old (VI) and a novel genotype (VII). Archiv. Virol. 143: 49-64. 1998. 16. Mayo, M A. A summary of the changes recently approved by ICTV. Archiv Virol 147,1655 - 1656. 2002. 17. Meulemans, G., M Gonze, M C. Carlier, P. Petit, A. Bumy, and L. E Long. Evaluation of the use of monoclonal antibodies to hemagglutination and fusion glycoproteins of Newcastle disease virus for virus identification and strain differentiation purposes. Arch. Virol. 92:55-62. 1987. 18. Rott, R., and H.-D. Klenk. Molecular basis of infectivity and pathogenicity of Newcastle disease virus. In: Newcastle disease. D. J. Alexander, ed. Kluwer Academic Publishers, Boston, Mass. pp. 98-112. 1988. 19. Seal, B. S., D. J. King, and J. D. Bennett. Characterization of Newcastle disease virus isolates by reverse transcription PCR coupled to direct nucleotide sequencing and development of sequence database for pathotype prediction and molecular epidemiological analysis. J. Clin. Microb. 33(10):2624-2630. 1995. 20. Srinivasappa, G. B., D. B. Snyder, W. W. Marquardt, and D. J. King, Isolation of a monoclonal antibody with specificity for commonly employed vaccine strains of Newcastle disease virus. Avian Dis. 30:562-567. 1986. 21. Wise, M G., J. C. Pedersen, D. A. Senne, D. Kapczynski, D. J. King, D. L. Surarz, B. S. Seal, and E. Spackman. Development of a real time reverse transcription-polymerase chain reaction for detection of Newcastle disease virus RNA in clinical samples. J. Clin. Microbiol. 42(1): 329-338. 2004.

REFERENCES

1. Adair, B. Μ, M S. McNulty, D. Todd. T .J. Connor, and K. Bums. Quantitative estimation of Newcastle disease virus antibody levels in chickens and turkeys by ELISA. Avian Pathol. 18:175-192. 1989. 2. Aldous, E. W. and D. J. Alexander, Technical Review: Detection and differentiation of Newcastle disease virus (avian paramyxovirus type 1) Avian Pathol. 30(2): 117-128. 2001. 3. Aldous, E. W., M. S. Collins, A. McGoldrick, and D. J. Alexander. Rapid pathotyping of Newcastle disease virus (NDV) using fluorogenic probes in a PCR assay. Vet. Microbiology. 80:201-212. 2001. 4. Aldous, E. W., Mynn, J. K. Banks, J. and D. J. Alexander, (A molecular epidemiological study of avian paramyxovirus type 1 (Newcastle disease virus) isolates by phylogenetic analysis of a partial nucleotide sequence of the fiision protein gene. Avian Pathol. 32: 239-357. 2003. 5. Alexander, D. Avian Paramyxoviridae— recent developments. In: Proceedings of the 1st European Society for Veterinary Virology Congress, Liege. Vet. Microbiol. 23:103-114. 1990. 6. Alexander, D.J. Newcastle disease, Other Avian Paramyxoviruses and Pneumovirus infections: Newcastle disease. In: Diseases of Poultry. Y.M Saif [ed in chief] Iowa State University Press USA pp 64-87. 2003. 7. Alexander, D. J., R. J. Manveil, J. P. Lowings, K. M Frost, M S. Collins, P. H. Russell, and J. E. Smith. Antigenic diversity and similarities detected in avian paramyxovirus type 1 (Newcastle disease virus) isolates using monoclonal antibodies. Avian Pathol. 26:399-418. 1997. 8. Ballagi-Pordany, A., E. Wehmann, J. Herczeg, S. Belak, and B. Lomniczi. Identification and grouping of Newcastle disease virus strains by restriction site analysis of a region from the F gene. Archiv. Virol. 141: 243261. 1996. 9. Beard, C. W., and R. P. Hanson. Newcastle disease. In: Diseases of poultry, 8th edition. M S. Hofstad, H. J. Barnes, B. W. Calnek, W. M Reid, and H. W. Yoder, eds. Iowa State University Press, Ames, Iowa, pp. 452470. 1984. 10. Creelan, J. L., D. A. Graham, and S. J. McCullough. Detection and differentiation of pathogenicity of avian paramyxovirus serotype 1 from field cases using one-step reverse transcriptase-polymerase chain reaction. Avian Pathology 31:493-499. 2002.

141

31 AVIAN METAPNEUMOVIRUS Richard E. Gough and Janice C. Pedersen

SUMMARY. Avian metapneumovirus (aMPV)) is an upper respiratory tract infection primarily of turkeys and chickens caused by a virus belonging to the genus metapneumovirus . The disease can cause significant economic losses in turkey flocks, particularly when exacerbated by secondary pathogens. Mortality is variable but may exceed 80% in susceptible turkey poults. In turkey breeding flocks infection can result in serious egg production problems. In chickens infection may lead to mild respiratory signs, egg production problems and swollen head syndrome (SHS). Agent Identification. Virus detection and identification can be difficult unless samples are taken early in the course of the disease. Virus isolation in cell cultures, embryonated hen’s eggs, tracheal organ cultures and molecular methods have all been used successfully to detect aMPV but the degree of success will depend on the strain of virus. Electron microscopy, virus neutralization and molecular techniques are widely used to identify the virus. The use of monoclonal antibodies and molecular studies have revealed antigenic diversity between the isolates studied. Serologic Detection in the Host. Confirmation of infection is best obtained by serologic tests, particularly enzyme-linked immunosorbent assay (ELISA) methods. In many countries where the disease is endemic vaccination is practiced which may complicate interpretation of results.

infraorbital sinuses, the so-called swollen head syndrome (SHS), torticollis, incoordination and depression. Unlike subtype A & B, the U.S. strain (subtype C) has not been shown to naturally induce disease in chickens. Previous work has demonstrated that different strains of a MPV have a specific tropism for chickens or turkeys (7). Although there is evidence that aMPV can infect other species of birds, clinical signs associated with the virus have been rarely reported (12). Subtype D metapneumoviruses have been reported to occur in ducks and is associated with respiratory signs and egg production problems (28).

INTRODUCTION

Avian metapneumovirus (aMPV) previously referred to as avian pneumovirus (APV) and avian rhinotracheitis (ART) virus is an acute, highly contagious upper respiratory tract infection of turkeys and chickens. In turkeys the virus causes a disease known as turkey rhinotracheitis (TRT). The agent responsible for the disease has been confirmed as a member of the genus Metapneumovirus in the family Paramyxoviridae (24). Until recently, APV was thought to be the sole member of the Metapneumovirus genus but recent reports from many countries have indicated that similar viruses have been detected in humans associated with respiratoiy tract infection in children (21). Avian metapneumovirus (aMPV) has been further classified into four subtypes: A, B, C and D, based on nucleotide and deduced amino acid sequence data (10). It is probable that other subtypes exist but have not yet been detected and identified. Serologic evidence suggests that the disease is now widespread throughout the world and of considerable economic importance, particularly in turkeys. Australasia appearing to be the only region in which aMPV has not been reported. There is serological and molecular evidence that aMPV occurs in a variety of other avian species, including pheasants, guinea fowl, ostriches, passerines and various waterfowl (21), but there is no evidence that serious disease occurs in these species.

SAMPLE COLLECTION It is very important to take samples for attempted virus isolation in the early stages of infection. Ideally live birds in the acute phase of the disease should be sampled using sterile swabs from the upper respiratory tract. The samples for virus isolation have been nasal exudates, choanal cleft swabs and scrapings of sinus and turbinate tissue. The virus has also been isolated from trachea and lungs, and occasionally viscera of affected turkey poults. Isolation of virus is rarely successful from birds showing severe chronic signs as the extreme clinical signs are usually due to secondary adventitious agents. This certainly applies to SHS of chickens in which the characteristic signs appear to be due to secondary Escherichia coli infection. Furthermore, for reasons that are unclear, virus isolation from chickens appears to be more difficult than from turkeys. For samples requiring dispatch to a diagnostic laboratory, it essential that the samples are sent immediately on ice. Where delays of more than three days are expected, the samples should be frozen when collected and sent frozen. Swabs for attempted virus isolation should be sent on ice fully immersed in virus transport medium but those for PCR analysis can be sent dry. For virus isolation, a 20% (v/v) suspension of the nasal exudate or homogenised tissue is made in phosphate-buffered saline (PBS) at pH 7.0-7.4 containing antibiotics. This is then clarified by centrifugation at 1000 * g for 10 min and the supernatant passed through a 450 nm membrane filter.

CLINICAL DISEASE

The various clinical signs associated with aMPV infection in turkeys have been detailed (13). Signs in young turkey poults include snicking, rales, sneezing, nasal discharge, foaming conjunctivitis, swelling of the infraorbital sinuses and submandibular edema. Secondary adventitious agents can dramatically exacerbate the clinical signs. In laying turkeys, infection normally results in only mild respiratory signs but with an associated marked drop in egg production, as much as 70%. In some flocks of adult turkeys, subclinical infections have been detected by seroconversion. When disease is seen, the morbidity can be as high as 100%, with mortality ranging from 0.5% in adult birds to 85% in young poults. The clinical signs of infection in chickens are less characteristic than the disease in turkeys. In laying flocks, particularly broiler breeders, there is a marked drop in egg production, often preceded by respiratory signs. Severe respiratoiy distress may occur in broiler chickens particularly when exacerbated by secondary pathogens such as infectious bronchitis virus, mycoplasmas, and Escherichia coli. The clinical signs associated with this may be swelling of the periorbital and

PREFERRED CULTURE MEDIA AND SUBSTRATES

To maximize the chances of successfully isolating the virus, a multiple approach to diagnosis is recommended. This is particularly relevant when dealing with different subtypes or genotypes which may require varied in vitro methods to isolate the virus. This was illustrated particularly well in North America where it was shown that primary isolation of subtype C aMPV was not associated with ciliostasis in tracheal organ cultures. This was in contrast to the 142

Chapter 31

European experience and elsewhere in which tracheal organ cultures were shown to be the most reliable method for the primary isolation of subtype A and B aMPV (10).

Avian Metapneumovirus

is rapidly destroyed by lipid solvents, heat (56 C for 30 min), and extremes of pH. Studies with a subtype C strain of virus, isolated from turkeys in the USA, reported that the virus was resistant to pH 5-9 for one hour. The study also reported that the viability of the virus was significantly reduced after 6 hr at 50 C, 2 days at 37 C, 4 wk at 20 C and less than 12 wk at 4 C. In addition, the study reported that several disinfectants were effective in reducing the viability of the virus (29)

Tracheal Organ Culture Tracheal organ cultures are prepared from turkey embryos or very young turkeys obtained from flocks free of specific antibodies to aMPV. Tracheas from chicken embryo or l-to-2-day-old chicks may also be used. Transverse sections of trachea are rinsed in PBS (pH 7.2), placed one to a tube in Eagles medium with antibiotics, and held at 37 C. For inoculation with infective material, the tubes are drained, and 0.1 ml of bacteria-free inoculum is added. After incubation for 1 hr at 37 C, growth medium is added and the cultures are incubated at 37 C on a roller apparatus rotating at 30 revolutions per hour. Cultures are examined daily after agitation on a laboratory mixer to remove debris from the lumen. Ciliostasis may occur within 7 days of inoculation on primary passage but usually is produced rapidly and consistently only after several blind passages.

Biological Properties Unlike other members of the family Paramyxoviridae, aMPV do not possess hemagglutination (HA) or neuraminidase activity.

Molecular Identification Due to the fastidious nature of aMPV, RT-PCR is significantly more sensitive and rapid method for the detection of aMPV than standard virus isolation methods (1,23). RT-PCR procedures targeted to the F, M, and G genes are used for the detection of aMPV, but are limited in specificity and have not been shown to detect all subtypes (3,4,22,23). These subtype specific assays are successfully used for the detection and diagnosis of endemic strains (17,23,26). However, limitations of subtype specific assays need to be recognized when conducting diagnostic testing for respiratory disease. Primers directed to conserved regions of the N gene have been shown to have broader specificity, detecting representative isolates from A, B, C, and D subtypes (3). RT-PCR assays directed to the G gene have been successfully used for genotype or subtype identification (14,15,17). Nasal exudates, choanal cleft swabs, and turbinate specimens collected 2-7 days post exposure are the preferred specimen. It has been shown, that aMPV can be detected from specimens collected 7-10 days post exposure, however, the viral concentration is considerably less thus reducing the success of detection (1,23). Five swabs from a single flock can be pooled to increase recovery rate. Isolation of aMPV from chickens is difficult and has succeeded only in a limited number of cases, for this reason, molecular tests are the method of choice for the detection of aMPV in chickens (16). In addition, RT-PCR has been used to genotype and confirm the identity of an aMPV isolated from chickens (17,27). It is important to remember that PCR detects viral RNA, not live virus; therefore, a positive PCR does not necessarily confirm an active infection.

Culture in Embryonating Eggs Six to 8-day-old embryonating chicken or turkey eggs, from flocks known to be free of aMPV antibodies, are inoculated by the yolksac route with 0.1-0.2 ml of bacteria-free material from infected birds and incubated at 37 C. Within 7-10 days, there is usually evidence of stunting of the embryos with few deaths. Consistent embryo mortality is normally seen only after four to five passages so this method of isolation is both time consuming and expensive.

Cell Culture Various cell cultures have been used for the primary isolation of aMPV, including chicken embryo cells, VERO cells and more recently the QT-35 cells, with varying degrees of success. However, once the virus has been adapted to growth in embryonating eggs or tracheal organ cultures, in which it grows only to low titers, the virus will readily replicate to high titers following multiple passages in a variety of primary chicken or turkey embryo cells and in Vero, BSC-1 and QT-35. The virus produces a characteristic cytopathic effect (CPE) with syncytial formation within 7 days. AGENT IDENTIFICATION Morphology By negative-contrast electron microscopy, the virus can be identified as having a paramyxovirus-like morphology. Pleomorphic fringed particles, roughly spherical and 80-200 nm in diameter, are commonly seen. Occasionally much larger filamentous forms are present, which may be up to 1000 nm in length. The surface projections are 13-14 nm in length and the helical nucleocapsid, that can sometimes be seen emerging from disrupted particles, is 14 nm in diameter with an estimated pitch of 7 nm per turn.

Antigen Detection A number of different assays have been developed for the detection of aMPV antigens using immunostaining methods. The most popular techniques have involved the use of immunoperoxidase (IP), immunofluorescence (IF) and immunogold staining and are described in detail elsewhere (10). These techniques, in particular IP and IF, have been used to detect virus specific antigen in both fixed and unfixed tissues and smears from turkeys and chickens. Immunological Detection Techniques Monoclonal antibodies have been used in virus neutralization tests to differentiate subtypes of aMPV (6,8). Neutralization tests using monospecific antiserum can also be used to confirm the identity of viruses isolated in cell or organ cultures. Because of low concentrations of virus, the method most commonly used is the alpha test (constant serum-diluted virus). The serum-virus mixtures are incubated at 37 C for 45 min and then assayed in cell or organ cultures for viable virus. A reduction in infectivity titer of 102.0 or more is considered to be significant. The immunodiffusion test has also been used to confirm the identity aMPV isolates (13). Briefly, isolates are cultivated in cell cultures to give an infectivity titer of approximately 106.0 median tissue culture infective doses (TCID50) per ml. After extensive CPE

Physiochemical Properties The genome of aMPV is unsegmented and composed of single­ stranded negative sense RNA of approximately 15 kilobases with a helical symmetry. In sucrose gradients, the buoyant density of an isolate from turkeys was found to be 1.21 g/ml with an approximate molecular weight of 500 x 106. The same virus was also shown to have 8 structural polypeptides of which 2 were glycosylated and 3 were non-structural virus-specified proteins. These have now been identified as follows; nucleoprotein (N), phosphoprotein (P), matrix protein (M), second matrix protein (M2), surface glycoprotein (G), fusion protein (F), a small hydrophobic protein (SH) and a viral RNA-dependant RNA polymerase (L). In common with other members of the family Paramyxoviridae, the infectivity of the virus 143

Richard E. Gough and Janice C. Pedersen

has occurred, the cell debris is removed and the supernatant concentrated by ultracentrifugation. The resulting concentrates are treated with 0.2% N-lauroylsarcosine (Sigma, St. Louis, Mo.) to produce antigens for the immunodiffusion test. Using monospecific aMPV antiserum, the antigens are tested by double immunodiffusion in 1% agarose. After approximately 24 hr at 37 C, the tests are examined and any precipitin lines that stain with 02% Coomassie brilliant blue R are interpreted as positive. Included in the test are appropriate positive and negative control antigens and sera. A precipitin line between the reference monospecific antiserum and test antigen confirms the identity of the virus.

The VN test can be carried out in confluent monolayers of chicken embryo cells in 96-well flat-bottomed microtitre plates. Briefly, 30300 median TCID of virus are reacted with two-fold serial dilutions of test serum at 37 C in 5% CO2. After 1 hr, 25ul is transferred from each dilution in the neutralization plate to the corresponding well on the cell culture plate. The plates are sealed and incubated at 37 C in 5% CO2, together with appropriate serum and virus controls. After 1 hr, 200 μΐ of maintenance medium is added to each well, the plates are sealed and incubated at 37 C in CO2 for 7 days. The cultures are examined daily for CPE and after 7 days the serum titres are determined and expressed as the reciprocal (log2) of the highest dilution of serum that completely inhibits viral CPE. A titre of >23 is considered positive.

SEROLOGIC DETECTION IN THE HOST

DIFFERENTIATION FROM CLOSELY RELATED AGENTS Due to difficulties in isolating and identifying aMPV, confirmation of infection is usually achieved by serological methods, particularly in unvaccinated chicken flocks. The most commonly employed method is the enzyme-linked immunosorbent assay (ELISA). Other methods that have been used to detect aMPV antibodies are virus neutralization, microimmunofluorescence and immunodiffusion tests. Ideally, both acute and convalescent serum samples should be obtained for testing. The sera should be heated at 56 C for 30 min before testing; if the testing of the sera is delayed, it should be stored at -20 C. In chickens, the serological response to aMPV infection is weak when compared to the response in turkeys (7).

Strain Variability When aMPV was first detected in Europe, it was believed there was only one serotype of the virus, represented by subtypes A and B. These were differentiated on the basis of nucleotide sequence analysis of the attachment (G) protein gene (14) and by mAb analyses (6,8). However, this situation changed with the emergence of a different aMPV in North America, designated subtype C (9,25). More recently, reports from France have indicated the presence of a fourth subtype of aMPV, designated subtype D (2,28). It is probable that other strains of aMPV exist, which are genetically distinct from subtypes A, B, C and D. These other subtypes of the aMPV may remain undetected using conventional f*CR based techniques; therefore a multi diagnostic approach is recommended when investigating outbreaks of respiratory disease in poultry.

Enzyme-Linked Immunosorbent Assay Numerous commercial ELISA kits, together with in-house assays, have been developed for the detection of aMPV antibodies (5,10) Although the indirect ELISA has been very useful for screening large numbers of sera for aMPV antibodies, differences in sensitivity and specificity between tests have been reported to occur between commercial kits (18,19). This is principally due to variations in the antigenicity and purity of the viral antigen used in the preparation of the ELISA kit. It appears that that sensitivity of the ELISA is less when a heterologous strain of aMPV is used as antigen, even though the strain appears closely related by virus neutralization test (10) Competitive or blocking ELISA kits have also been developed, incorporating an aMPV specific monoclonal antibody (mAb).These kits claim to have a broad spectrum of sensitivity and specificity for all subtypes of aMPV and can be used for testing sera from a variety of avian species. ELISA antigens have been prepared in a variety of substrates including various cell cultures and tracheal organ cultures (10).

Newcastle Disease Some strains of Newcastle disease virus and other members of the genus Paramyxovirus, such as PMV-3 (see Chapter 30 on Newcastle disease virus and other avian paramyxoviruses), may cause respiratory disease and egg production problems in chickens and turkeys that closely resemble aMPV. Paramyxoviruses are similar in morphology but can usually be easily distinguished from aMPV because they possess HA and neuraminidase activity.

Infectious Bronchitis Infections of chickens with infectious bronchitis (IB) virus can result in respiratory disease and egg production problems that are similar to aMPV. Swollen head syndrome (SHS) in chickens has also been described as being associated with infectious bronchitis virus and E. coli (11,20) Serologic and molecular identification of the infecting virus is probably the simplest method of making a differential diagnosis.

Fluorescent Antibody Test Several techniques have been described for the detection of aMPV antibodies using indirect immunofluorescence (HF) tests on infected tissues or cell cultures. Studies have shown that antibodies to aMPV in infected turkeys can be detected by HF 5 days after the appearance of clinical signs (10). The technique has limited application for the large scale testing of poultry sera and has been mainly used for research purposes.

Avian Influenza Infection of chickens or turkeys with the milder strains of avian influenza viruses can result in disease similar to aMPV. Virus isolation and the demonstration of HA activity by influenza viruses will distinguish between the viruses.

Bacteria and Mycoplasma A wide range of bacteria and Mycoplasma species have been reported to cause disease signs very similar to aMPV. Frequently, such organisms may be present as secondary or adventitious invaders and may cause considerable diagnostic problems. Distinction must rely on negative isolation or demonstration of antibodies to aMPV .

Virus Neutralization Test Antibodies to the virus have been detected by standard virus neutralization (VN) techniques in tracheal organ cultures and various sensitive cell cultures, such as CEF, chicken embryo liver, and VERO cultures (13). Using a neutralization assay in CEF cells, it was shown that neutralizing antibody could be detected within 5 days of the appearance of clinical signs and was declining by day 13. There is a good correlation between VN results and ELISA and immunofluorescence results (10). There are cross reactions between subtype A and B viruses so the VN test is not suitable for distinguishing subtype antibodies.

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Avian Metapneumovina

REFERENCES

1. Alkhalaf, A.N., L.A.Ward, RN.Dearth and Y. M Saif. Pathogenicity, transmissibility and tissue distribution of avian pneumovirus in turkey poults. Avian Dis. 46:650-659. 2002. 2. Bayon-Auboyer, Μ H., C. Amauld, D.Toquin and N. Eterradossi. Nucleotide sequences of the F, L and G protein genes of two non-A/non-B avian pneumoviruses (APV) reveal a novel APV subgroup. J. Gen. Virol. 81:2723-2733. 2000. 3. Bayon-Auboyer, MH., V. Jestin, D. Toquin, M Cherbonnel and N. Eterradossi. Comparison of F-, G- and N-based RT-PCR protocols with conventional virological procedures for the detection and typing of turkey rhinotracheitis virus. Arch Virol. 144:1091-1109. 1999. 4. Bennett, R.S., J. Nezworski, B.T. Velayudhan, K.V. Nagaraja, D. H.Zeman, N.Dyer, T.Graham, D.C. Lauer, MK. Njenga and D.A. Halvorson. Evidence of avian pneumovirus spread beyond Minnesota among wild and domestic birds in central North America. Avian Dis. 48:902-908. 2004. 5. Chiang, S.J., A.M Dar, S.M Goyal, K.V. Nagaraja, D.A. Halvorson and V. Kapur. A modified enzyme-linked immunosorbent assay for the detection of avian pneumovirus antibodies. J. Vet. Diagn. Invest. 12:381-384. 2000. 6. Collins, MS., R .E .Gough and D .J .Alexander. Antigenic differentiation of avian pneumovirus isolates using polyclonal antibody and mouse monoclonal antibodies. Avian Pathol. 22:469-479. 1993. 7. Cooke, J.K.A., S. Kinloch and MM. Ellis. In vitro and in vivo studies in chickens and turkeys on strains of turkey rhinotracheitis virus isolated from the two species. Avian Pathol. 22:157-170. 1993a. 8. Cook, J. K. A., Β. V. Jones, Μ. M Ellis, Jing Li and D. Cavanagh. Antigenic differentiation of strains of turkey rhinotracheitis virus using monoclonal antibodies. Avian Pathol. 22:257-273. 1993b. 9. Cook, J.K.A., MB. Huggins, S.J. Orbell and D.A. Senne. Preliminary antigenic characterization of an avian pneumovirus isolated from commercial turkeys in Colorado, USA. Avian Pathol. 28:607-617. 1999. 10. Cook, J.K.A. and D. Cavanagh. Detection and differentiation of avian pneumoviruses (metapneumoviruses). Avian Pathol. 31:117-132. 2002. 11. Droual, R. and P R. Woolcock. Swollen head syndrome associated with E. Coli and infectious bronchitis virus in the Central Valley of California. Avian Pathol. 23:733-742. 1994. 12. Gough, R.E., MS. Collins, W.J. Cox and N.J. Chettle. Experimental infection of turkeys, chickens, ducks, guinea fowl, pheasants and pigeons with turkey rhinotracheitis virus. Vet. Record. 123:58-59. 1988. 13. Gough, R.E. Avian pneumoviruses. In: Diseases of Poultry, 11th ed. Y. M Saif, H. J. Bames, J.R. Glisson, A M Fadly, L.R. McDougald and D.E. Swayne. Iowa State University Press, Ames, Iowa, pp 92-99. 2003. 14. Juhasz, K., and A. J. Easton. Extensive sequence variation in the attachment (G) protein gene of avian pneumovirus: evidence for two distinct subgroups. J. Gen. Virol. 75:2873-2880. 1994.

15. Lwamba, H.C.M, R. Alvarez, M.G. Wise, Qingzhong, Yu, D. Halvorson, MK. Njenga and B.S. Seal. Comparison of full-length genome sequence of Avian metapneumovirus subtype C with other paramyxoviruses^ Virus Research. 107:83-92. 2005. 16. Mase, M,S. Asahi, K.Imai, K.Nakamura and S. Yamaguchi. Detection of turkey rhinotracheitis virus from chickens with swollen head syndrome by reverse transcriptase-polymerase chain reaction (RT-PCR). J. Vet Med. Sci. 58:359-361.1996. 17. Mase, M, S. Yamaguchi, K. Tsukamoto, T. Imada, K. Imai and K Nakamura. Presence of avian pneumovirus subtypes A and B in Japan. Avian Dis. 47:481-484.2003. 18. McFarlane-Toms, I.P and R.J.H. Jackson. A comparison of three commercially available ELISA tests for detecting antibodies to turkey rhinotracheitis virus (TRTV). In: E. Kaleta and U. Heffels-Redman(Eds). Proceedings of the International Symposium on Infectious Bronchitis and Pneumovirus infections of Poultry. Rauischolzhausen, Germany, pp26-37. 1998. 19. Mekkes, D. R. and J. J. de Wit. Comparison of three commercial ELISA kits for the detection of turkey rhinotracheitis virus antibodies. Avian Pathol. 27: 301-305. 1999. 20. Morley, A.J. and D.K. Thomson. Swollen-head syndrome in broiler chickens. Avian Dis. 28:238-243. 1984. 21. Njenga, MK., H.M Lwamba and B.S. Seal. Metapneumoviruses in birds and humans. Virus Research. 91:163-169. 2003. 22.. Pedersen, J.C., L. Roland, D.L. Reynolds and A. Ali. The sensitivity and specificity of a reverse transcriptase-polymerase chain reaction assay for the βλάβη pneumovirus (Colorado strain). Avian Dis. 44:681-685. 2000. 23. Pedersen, J.C., D.A. Senne, B. Panigrahy and D.L. Reynolds. Detection of avian pneumovirus in tissue and swab specimens from infected turkeys. Avian Dis. 45:581-592. 2001 24. Pringle, C.R. Virus Taxonomy-San Diego. Arch.of Virology. 143:14491459. 1998. 25. Senne, D.A., RK. Edson., J.C. Pedersen and B. Panigrahy. Avian pneumoxdrus update. Proceedings of American Veterinary Medical Association 134th Annual Congress. Reno, NV, USA. ppi90.1997. 26. Shin, H.J., F.F. Jirjis, M.C. Kumar, MK. Njenga, D.P. Shaw, S.L. Noll, K.V.Nagaraja and D.A. Halvorsen. Neonatal avian pneumovirus infection in commercial turkeys. Avian Dis. 46:239-244. 2002. 27. Tanaka, M,H. Tanuma, N.Kokumai, E.Oishi, T. Obi, K. Hiramatsu and Y. Shimizu. Turkey rhinotracheitis virus isolated from broiler chicken with swollen head syndrome in Japan. J. Vet. Med. Sci. 57:939-945.1995. 28. Toquin, D., MH. Bayon-Auboyer, N. Etteradossi, H. Morin and V. Jestin. Isolation of a pneumovirus from a Muscovy duck. Vet. Record. 145:680 29. Townsend, E., D.A. Halvorson, K.E.Nagaraja and D.P.Shaw. Susceptibility of an avian pneumovirus isolated from Minnesota turkeys to physical and chemical agents. Avian Dis. 44:336-342. 2000.

32 INFECTIOUS BRONCHITIS Jack Gelb, Jr. and Mark W. Jackwood

SUMMARY. Infectious bronchitis (IB) is caused by infectious bronchitis virus (IBV), a member of the family Coronaviridae. IBV is highly host specific, causing natural infections mainly in chickens. Primary infections in young chickens typically produce respiratory disease but some strains may cause kidney lesions. Layers and breeders commonly suffer egg production losses with or without evidence of respiratory disease signs. Numerous serotypes of the virus have been responsible for outbreaks in commercial chickens in spite of the use of attenuated live and inactivated vaccines. IB is a growing problem for poultry producers because only a limited number of serotypes are available for vaccination, and the cross-protection elicited against unrelated field strains may be minimal. Agent Identification. Presumptive diagnosis is made in a suspect flock based on clinical signs consistent with the disease and evidence of IBV serum antibody production as demonstrated by enzyme-linked immunosorbent assay (ELISA). Isolation and identification of the causative serotype of IBV is required for definitive diagnosis. Serotype identification may be achieved by virus-neutralization, hemagglutination-inhibition, type-specific monoclonal antibodies, or reverse transcriptase-polymerase chain reaction followed by restriction fragment length polymorphism or sequencing. Cross-challenge tests in chickens may be used to determine the potential protection afforded by immunization with various IBV vaccines or field strains to control the disease. Serologic Detection in the Host. ELISA is preferred method for detecting serum antibody response, but it does not identify serotype­ specific antibodies. Acute and convalescent sera response is useful in sera diagnosis and for monitoring flock vaccination titers.

INTRODUCTION

SAMPLE COLLECTION

Infectious bronchitis (IB) is a highly contagious clinically acute disease of the respiratory and urogenital tract of chickens, caused by infectious bronchitis virus (IBV), a member of the family Coronaviridae. The disease is common throughout the world where chickens are produced commercially. Mixed infections involving IBV, Newcastle disease virus (NDV), avian adenovirus group 1, Mycoplasma, and coliform bacteria are common and may confuse diagnostic efforts. Many serotypes of IBV are recognized and have practical significance in the control of IB, because immunity following infection or vaccination with one serotype often is not protective against infections with unrelated serotypes. In the United States, the most often isolated IBV serotypes in commercial poultry are Arkansas (Ark), Connecticut (Conn), Delaware (DE/072/92), and Massachusetts (Mass). Previously unrecognized antigenic variants have been recovered from multiage commercial layer complexes with respiratory disease or egg production problems (10). Many antigenic variant serotypes also have been reported in European countries and Australia (7). Undoubtedly, many more strains will be isolated from these and other countries as emphasis on IBV surveillance increases. For an in-depth review of IB, refer to Cavanagh and Naqi (4). A detailed discussion of IBV antigen, genome and antibody detection assays prepared by De Wit (6) is also available.

Samples for IBV isolation must be obtained as soon as clinical disease signs are evident. Tracheal swabs are preferred and are placed directly into cold media with antibiotics to suppress bacterial and fungal growth and preserve the viability of the virus. Sterile swabs are used to swab 5-10 clinically affected birds per flock. Cotton-tipped swabs of varying sizes * are available (Fisher Scientific, Pittsburgh, Penn.) depending on the age and breed of the chicken. The swabs are placed in 2-3 ml of cold sterile tryptose phosphate broth (TPB) (pH 7.0-7.2) containing 10,000 IU/ml penicillin, 10,000 IU/ml streptomycin, and 250 IU/ml amphotericin B. Swab tubes are immediately placed on ice and are frozen at -20 C at the earliest convenience. Other tissues, such as lung, kidney, oviduct, cecal tonsil and proventriculus may be collected using aseptic techniques. Tissues are placed in clean, labeled, tightly sealed plastic specimen bags or sterile tubes and are frozen at -20 C. Cloacal swabs may be obtained and are handled as described above. All swabs and tissue samples should be frozen and transported in an insulated container to a diagnostic virology laboratory. Because IBV persists in the intestinal tract, isolation of some strains from cecal tonsil and cloacal swabs is possible for several weeks after the disappearance of clinical signs. Accordingly, isolating IBV from these sites does not confirm its role as the causative agent in a recent disease outbreak. The frozen TPB containing tracheal and cloacal swabs is thawed, mixed, and incubated at room temperature (about 22 C) for 30-60 min prior to inoculation to reduce the possibility of bacterial and fungal contamination. Contamination sometimes associated with virus isolation from cloacal swabs may be avoided by centrifuging the swab tube broth at 1000 x g for 15 min and then passing the supernatant fluid through a 0.22 or 0.45-μιη sterile syringe filter. Tissue homogenates (10% w/v) are prepared in TPB with antibiotics by disrupting lung, kidney, oviduct, or cecal tonsil using a glass tissue grinder (Tenbroeck, VWR Scientific Products, West Chester, Penn.) or mortar and pestle. The homogenates are incubated at room temperature for 30-60 min. Sentinel chickens have been used to help facilitate IBV isolation in commercial flocks (9). Specific-pathogen-free (SPF) chickens immunized against IBV vaccine serotype(s) are placed with IBVsusceptible sentinels in pens or cages in commercial chicken houses. After a 1-wk exposure period, the field-exposed sentinels are removed, and tracheal swabs are collected for virus isolation attempts. Several successive weekly placements increase the possibility of isolating IBV. The use of IBV-immune sentinels

CLINICAL DISEASE

Chickens are the primary natural host, although pheasants may be susceptible to IBV or highly similar coronaviruses (3). In chickens a short incubation period (24—72 hr) is a unique characteristic of the disease. Young chicks display acute respiratory disease signs and lesions in the trachea. Morbidity can approach 100%, but mortality is generally below 5% in outbreaks not complicated by secondary pathogens or concurrent infections. In layer chickens, IBV causes decreased egg production and quality. Lymphoid cell infiltration and epithelial cell degeneration of the oviduct wall have been observed. Nephropathogenic strains such as Holte, Gray, and Australian T produce enlarged kidneys with distended tubules and ureters containing uric acid crystals. Diarrhea, dehydration, depression, and death may occur in affected birds.

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provides a better opportunity for isolating IBV antigenic variants free of contaminating vaccine virus serotypes that may be cycling in commercial chickens.

Infectious Brondnta

DNA. Most strains of IBV are inactivated after 15 min at 56 C. IBV is inactivated by lipid solvents, such as chloroform and ether. Sensitivity to lipid solvents can also be used to screen LBV field isolates for contaminating naked viruses, such as avian adenovirus group 1 and reovirus.

PREFERRED CULTURE MEDIA AND SUBSTRATES

Infectious bronchitis virus is most commonly grown in embryonated eggs, tracheal organ culture (TOC), and chicken kidney cell culture. The embryonated egg is preferred for primary isolation attempts.

Serological Identification by Virus Neutralization (VN) and Hemagglutination-Inhibition (HI) Virus Neutralization. Neutralization of an IBV field isolate by known serotype-specific IBV antiserum establishes its identity. The VN test may be performed in embryonated eggs, chicken kidney cell culture, or TOC. The test may be conducted using the constant­ serum diluted-virus (alpha) or diluted-serum constant-virus (beta) method. In addition, a constant-virus constant-serum VN procedure (5) has been useful for serotyping IBV field isolates. VN tests are performed in embryonated eggs by reacting 32-320 mean embryo infectious doses (ΕΠ)50) of an IBV field isolate with antisera to known strains containing 1-20 units of antibody. The antibody unit concentration is determined by titration against the homologous IBV serotype. The highest dilution of antiserum protecting 50% of the inoculated embryos is equal to one antibody unit. As concurrent infections with two or more IBV serotypes are common, reciprocal VN tests are sometimes required to identify all IBV in field isolates. Antiserum to the field isolate is reacted against known IBV serotypes to establish the identities of the causative serotypes. Hemagglutination Inhibition. Hemagglutination antigen for the HI test is prepared from chorioallantoic fluid harvested from IBVinoculated embryonated eggs. Neuraminidase type V in PBS (pH 7.2) at 1.0 units/ml final concentration is used to treat IBV for 30 min at 37 C (24). HA antigen titration is performed in standard Ubottom 96-well microtiter plates using CRBCs. Treatment of IBV with bacterial phospholipase C was initially thought to enable the virus to agglutinate CRBCs (1,16). However, HA antigens produced using highly purified phospholipase C preparations often had considerably lower titers than those produced using unpurified phospholipase C preparations. Subsequent studies (24,25) determined that treatment of IBV with purified neuraminidase preparations consistently produced high-titer HA antigens. These findings suggested that the unpurified phospholipase C preparations were contaminated with neuraminidase. The microtiter HI test may be used to serotype IBV (1,16). Neuraminidase-treated HA antigen is first prepared from the field isolate. A concentration of 4-8 HA units of antigen is added to twofold dilutions (1:2 to 1:1024) of serotype-specific known IBV antiserum. After incubation at room temperature for 30 min, CRBCs (0.5%-1.0%) are added, and the test is read 30-60 min later. The inhibition of HA of the field isolate by a known antiserum identifies the serotype of IBV but caution must be used when interpreting this test because cross-reactivity is not uncommon. Production of Serotyping Antiserum for VN and HL Antiserum suitable for serotyping IBV isolates using the VN or HI procedures is produced preferably in SPF chickens. Typically, 3-to-8-wk-old chickens are inoculated intratracheally with about 105 ELD50 of IBV per bird. The chickens are placed in isolation rooms or Horsfall isolation units. Two weeks PI, the chickens are reinoculated by intravenous injection with at least ΙΟ5 ΕΠ)50 of IBV. Blood samples are collected from chickens 2 wk after intravenous injection of IBV. Serum is harvested, pooled, and inactivated at 56 C for 30 min before being used in VN or HI serotyping procedures. It is important for antisera to be highly specific for the homologous IBV serotype and to not cross-react with heterologous virus serotypes. Repeated inoculations (three or more) or the use of adjuvants in preparing serotyping antisera is not advisable because antibodies may cross-react with heterologous IBV serotypes.

Embryonated Chicken Eggs Embryonated chicken eggs from an SPF source are recommended for virus isolation. Ten 9 to 11-day-old embryos are inoculated by the chorioallantoic sac route with 0.2 ml of swab tube broth or tissue homogenate. The eggs are candled daily, with mortality between days 2 and 7 postinoculation (PI) considered to be virus­ specific. On day 3 PI, five eggs are removed from the incubator and are placed at 4 C for 18-24 hr. Chorioallantoic fluid harvested aseptically from inoculated eggs should be free of bacteria and fungi and give a negative hemagglutination (HA) reaction with chicken red blood cells (CRBCs). The remaining embryos are incubated up to 7 days and are then observed for typical IBV lesions, such as stunting, curling, clubbing of the down, or urate deposits in the mesonephros of the kidney. Cutaneous hemorrhage is often noted with embryo-lethal strains of IBV. Chorioallantoic membranes are collected, homogenized, and tested for avian adenovirus group 1 by the immunodiffusion method. Group 1 adenovirus infections of commercial chickens are common, and the virus often produces stunted embryos indistinguishable from IBVinfected embryos. Some IBV field isolates are not embryo-adapted and do not cause death or produce lesions on the first passage. Therefore, a minimum of three passages should be made before a virus isolation attempt is considered to be negative. Particular attention should be given to urate deposits in the mesonephros of the kidney in evaluating embryos inoculated with field isolates.

Organ Cultures Chicken TOC may be used to isolate IBV. The primary advantage in using TOC over embryonated eggs is that non-embryo-adapted field strains of IBV produce a rapid ciliostasis in TOC on the first passage, eliminating the need for multiple passages. A disadvantage is that a cell culture facility is needed for preparing and maintaining TOC. Other potential limitations of TOC involve differentiating or detecting viruses other than IBV in field samples. NDV and IBV produce extensive ciliostasis by 3 days PI. Avian adenovirus replicates in TOC, but because it does not produce rapid and complete ciliostasis, its presence may go undetected.

Cell Cultures Primary isolation of IBV in chicken kidney cell culture is not recommended, because the virus requires adaptation in embryonated eggs before it will grow in cell culture. AGENT IDENTIFICATION

Physicochemical Properties Coronaviruses are enveloped, single-stranded RNA viruses with helical symmetry (see Chapter 45 on virus identification and classification for characterization techniques). The virion is pleomorphic and has a diameter of 80-120 nm. The envelope consists of large club-shaped spike proteins that give the surface of the virion its characteristic corona appearance. Coronavirus replication occurs in the cytoplasm. Inhibitors of DNA, such as the halogenated nucleotide 5-iodo-2-deoxyuridine, do not inhibit IBV multiplication, indicating that the virus contains RNA and not 147

Jack Gelb, Jr. and Mark W. Jackwood

serotypes may produce a greater degree of protection than would be predicted on the basis of serotyping results. Cross-challenge tests are conducted by immunizing ten 3-to-6-wkold SPF chickens by eye drop with known IBV serotypes and challenging them 4 wk later by eye drop inoculation with an unknown field isolate. Appropriate controls are included in cross­ challenge tests. The immunized chickens challenged with the homologous strain must be protected. Nonimmunized control chickens inoculated with the challenge virus must be susceptible. Isolation facilities are required to prevent inadvertent cross­ infection between groups of chickens immunized with different serotypes. Protection is evaluated most commonly by collecting tracheal swabs at 4 or 5 days after challenge and assessing them for challenge virus by performing virus isolation attempts in embryonated eggs. Chickens from which virus is not isolated are considered to be protected. Other methods for determining protection following challenge include evaluating microscopic lesions in the trachea or measuring ciliary activity in TOC explants (20). Antigenically novel field isolates should be used as challenge viruses following immunization of SPF chickens with vaccine(s) containing the serotype(s) currently approved for use in the region or country where the isolate was recovered. The results of these studies will establish the cross-protective potential of a vaccine(s) against field isolate challenge.

Monoclonal Antibody Identification of IBV Antigens Type-specific monoclonal antibodies specific to the SI subunit of the spike glycoprotein of serotypes Mass, Conn, and Ark have been produced (13) and used to identify the respective serotypes by antigen-capture enzyme-linked immunosorbent assay (AC-ELISA) (23). Other serotypes may be identified as IBV but not identified to the specific serotype using a group-specific monoclonal to the M glycoprotein of IBV (13, 23). Allantoic fluid from IBV-inoculated embryonated chicken eggs is best suited for identification of IBV by AC-ELISA although tissue homogenates can be used. Immunoperoxidase staining (22) and immunofluorescent antibody assays have also been used to identify serotypes using type-specific IBV monoclonal antibodies. Molecular Identification Several reverse transcriptase-polymerase chain reaction (RT-PCR) assays have been developed for identifying IBV serotypes. The assays use serotype-associated sequence variations in the SI subunit of the spike glycoprotein gene to identify and differentiate strains and variants. Generally, allantoic fluids harvested following the inoculation of eggs with clinical samples are used for RNA extraction and RTPCR amplification (2, 11, 12). A RT-PCR restriction fragment length polymorphism (RFLP) assay has been developed that differentiates IBV types based on unique electrophoresis banding patterns of restriction enzyme-digested fragments of the RT-PCR amplified SI gene (12,18). The RT-PCR RFLP test can identify all known serotypes of IBV as well as variant viruses. The RT-PCR RFLP procedure may be used in conjunction with a RT-PCR assay that uses universal primers to the conserved membrane and nucleocapsid genes (2). Although that assay does not identify virus type, it does amplify all types of IBV and can be used in conjunction with a biotin-labeled DNA probe (11), or nucleic acid sequencing to verify the presence of IBV in the sample SI genotype-specific RT-PCR can also be used to identify specific IBV serotypes (14). SI gene primers specific for serotypes Mass, Conn, Ark, and JMK are used in conjunction with a universal primer set that amplifies all IBV serotypes. Specific primers for the DE/072/92 and California serotypes have also been developed. Other IBV types can be detected using the general primers, but the specific serotype cannot be identified. Both RT-PCR RFLP and genotype specific RT-PCR are capable of detecting more than one IBV type in a clinical sample. In addition, clinical samples inactivated with an equal volume of buffered (pH 4.5) phenol or spotted on Finders Technology Associates (FTA) cards (Whatman Inc. Florham Park, NJ) can be used as a source of template for RTPCR amplification (21). These inactivated samples can be safely transported to the laboratory and with the proper permits they can be imported from abroad. Cycle sequencing of the RT-PCR amplified hypervariable amino terminus region of SI may be used to identify previously recognized field isolates and variants (17,19). Comparison and analysis of sequences of unknown field isolates and variants with reference strains available in GenBank (National Center for Biotechnology Information http://www.ncbimlm.nih.gov/) can establish potential relatedness and is a significant advantage of sequencing. The major uses of RT-PCR tests are virus identification and its application in the understanding of epidemiological investigations during IBV outbreaks. The RT-PCR tests, as they now exist however, do not provide information on viral pathogenicity.

SEROLOGIC IDENTIFICATION IN THE HOST

Serodiagnosis of IBV infection in commercial chickens is best performed by demonstrating an ascending serum antibody response in recovered chickens using the enzyme-linked immunosorbent assay (ELISA). Commercially available ELISA kits detect antibodies common to all IBV serotypes (IDEXX Laboratories, Inc., Westbrook, Maine; Synbiotics Corp., San Diego, California) and are not capable of identifying serotype-specific antibodies induced by the IBV strain(s) responsible for the outbreak. Nonetheless, the ELISA is a very valuable, inexpensive, and easy to use tool for serodiagnosis of IBV to rule out other common causes of respiratory disease such as NDV, infectious laryngotracheitis virus, or Mycoplasma. Commercial IBV ELISA kits with computerfacilitated data analysis capabilities are available. ELISA systems using 96-well microtiter plates and a single-serum dilution approach are used to evaluate immune status in commercial flocks. Acute and convalescent serum samples, preferably from the same chickens in a suspect IBV-infected flock, are tested by ELISA. It is recommended that serum from acute samples be stored at -20 C until convalescent samples are obtained. Both groups of sera can then be run at the same time to minimize variability. An increase in antibody titer between acute and convalescent serum samples is indicative of an IBV infection. Commercial ELISA systems for measuring IBV serum antibody have been compared with VN and HI serology and found to give favorable results (26,27). The use of VN or HI tests for serodiagnosis is generally not recommended. These tests are expensive and tedious to perform on a routine basis. In addition, interpretation of the results may be difficult because sera of commercial chickens often contain cross­ reacting antibodies resulting from multiple infections with heterologous vaccinal and field strain serotypes (8). Antibodies produced by young chickens tend to be more serotype-specific, whereas cross-reactions with heterologous VN and HI antigens are common in breeders and layers (15).

Cross-Challenge Studies Cross-challenge tests in chickens are an important adjunct to SI genotyping or serotyping for antigenically characterizing an IBV field isolate. Immunization of chickens with antigenically distinct 148

Chapter 32 Infectious Bronchitis 14. Keeler, C. L., K. L. Reed, W. A. Nix, and J. Gelb. Serotype identification of avian infectious bronchitis virus (IBV) by RT-PCR of the peplomer (S-l) gene. Avian Dis. 42:275-284. 1998. 15. King, D. J., and S. R. Hopkins. Evaluation of the hemagglutination­ inhibition test for measuring the response of chickens to avian infectious bronchitis virus vaccination. Avian Dis. 27:100-112. 1983. 16. King, D. J., and S. R. Hopkins. Rapid serotyping of infectious bronchitis virus isolates with the hemagglutination-inhibition test. Avian Dis. 28:727— 733. 1984. 17. Kingham, B. F., C. L. Keeler, Jr., W. A. Nix, B. S. Ladman, and J. Gelb, Jr. Identification of avian infectious bronchitis virus by direct automated cycle sequencing of the S-lgene. Avian Dis. 44:325-335. 2000. 18. Kwon, Η. Μ, M W. Jackwood, and J. Gelb, Jr. Differentiation of infectious bronchitis virus serotypes using polymerase chain reaction and restriction fragment length polymorphism analysis. Avian Dis. 37:194-202. 1993. 19. Lee, C-W., D. A. Hilt, and M. W. Jackwood. Typing of field isolates of infectious bronchitis virus based on the sequence of the hypervariable region in the SI gene. Veterinary Diagnostic Investigation. 15:344-348. 2003 20. Marquardt, W. W., S. K. Kadavil, and D. B. Snyder. Comparison of ciliary activity and virus recovery from tracheas of chickens and humoral immunity after inoculation with serotypes of avian infectious bronchitis virus. Avian Dis. 26:828-834. 1982. 21. Moscoso, Η., E. O. Raybon, S. G. Thayer, and C. L. Hofacre. Molecular detection and serotyping of infectious bronchitis virus from FTA filter paper. Avian Dis. 49:24-29. 2005. 22. Naqi, S. A monoclonal antibody-based immunoperoxidase procedure for rapid detection of infectious bronchitis virus in infected tissues. Avian Dis. 34:893-898. 1990. 23. Naqi, S. A., K. Karaca, and B. Bauman. A monoclonal antibody-based antigen capture enzyme-linked immunosorbent assay for identification of infectious bronchitis virus serotypes. Avian Pathol. 22:555-564. 1993. 24. Ruano, M, J. El-Attrache and P. Villegas. A rapid-plate hemagglutination assay for the detection of infectious bronchitis virus. Avian Dis. 44:99-104. 2000. 25. Shultze, B., D. Cavanagh, and G. Herrler. Neuraminidase treatment of avian infectious bronchitis coronavirus reveals a hemagglutinating activity that is dependent on sialic acid-containing receptors on erythrocytes. Virology 189:792-794. 1992. 26. Thayer, S. G., B. N. Nersessian, B. Rivetz, and O. J. Fletcher. Comparison of serological tests for antibodies against Newcastle disease virus and infectious bronchitis virus using ImmunoComb® solid-phase immunoassay, a commercial enzyme-linked immunosorbent assay, and the hemagglutination-inhibition assay. Arian Dis. 31:459-463. 1987. 27. Thayer, S. G., P. Villegas, and O. J. Fletcher. Comparison of two commercial enzyme-linked immunosorbent assays and conventional methods for avian serology. Arian Dis. 31:120-124. 1987.

DIFFERENTIATION FROM CLOSELY RELATED AGENTS

Respiratory disease associated with IBV infections may be clinically indistinguishable from mild respiratory forms of Newcastle disease, infectious laryngotracheitis, and low pathogenic avian influenza. In these cases, diagnosis depends on isolation and identification of the virus or demonstration of an increase in specific antibody production associated with recovery from infection. Virulent strains of NDV and infectious laryngotracheitis virus cause a more severe disease than occurs in IB outbreaks. Neurological signs or visceral lesions with high mortality are observed in flocks infected with virulent NDV. Infectious laryngotracheitis virus can produce a hemorrhagic tracheitis with high mortality in serious outbreaks. In addition, infectious laryngotracheitis spreads more slowly in affected flocks. Infectious coryza and mycoplasmosis may also resemble IB complicated by pathogenic coliform bacteria. Swollen head syndrome is associated with IBV, Escherichia coli, and high ammonia levels. Facial swelling is also observed in infectious coryza and Mycoplasma infections. Nephritis observed in chickens infected with nephropathogenic strains of IBV resembles kidney changes seen in several disease conditions, such as infectious bursal disease, mycotoxicosis, or other toxicities. REFERENCES

1. Alexander, D. J., and N. J. Chettle. Procedures for the haemagglutination and the haemagglutination inhibition tests for avian infectious bronchitis virus. Avian Pathol. 6:9-17. 1977. 2. Andreasen, J. R. Jr., M W. Jackwood, and D. A. Hilt. Polymerase chain reaction amplification of the genome of infectious bronchitis virus. Avian Dis. 35:216-220. 1991. 3. Cavanagh, D., P. Britton, R. E. Gough, K. Mawditt, D. de B. Welchman. Coronaviruses from pheasants (Phasianus colchicus) are genetically closely related to coronaviruses of domestic fowl (infectious bronchitis virus) and turkeys. Avian Pathol. 31:81-93. 2002. 4. Cavanagh, D. and S. A. Naqi. Infectious bronchitis. In: Diseases of poultry. 11“' ed. Saif, Y. Μ., H. J. Bames, J. R. Glisson, A. M. Fadly, L. R. McDougald, and D. E. Swayne. Iowa State University Press Ames, LA. 1 Ol­ li 9. 2003. 5. Cowen, B. S., and S. B. Hitchner. Serotyping of avian infectious bronchitis viruses by the virus-neutralization test. Avian Dis. 19:583-595. 1975. 6. De Wit, J.J. Technical review. Detection of infectious bronchitis virus. Avian Pathol. 29:71-93. 2000. 7. Gelb, J., Jr., C. L. Keeler, Jr., W. A Nix, J. K. Rosenberger, and S. S. Cloud. Antigenic and S-l genomic characterization of the Delaware variant serotype of infectious bronchitis virus. Avian Dis. 41:661-669. 1997. 8. Gelb, J., Jr., and S. L. Killian. Serum antibody responses of chickens following sequential inoculations with different infectious bronchitis virus serotypes. Avian Dis. 31:513-522. 1987. 9. Gelb, J., Jr., J. K. Rosenberger, P. A. Fries, S. S. Cloud, E. M Odor, J. E. Dohms, and J. S. Jaeger. Protection afforded infectious bronchitis virusvaccinated sentinel chickens raised in a commercial environment. Avian Dis. 33:764-769. 1989. 10. Gelb, J., Jr., J. B. Wolff, and C. A. Moran. Variant serotypes of infectious bronchitis virus isolated from commercial layer and broiler chickens. Avian Dis. 35:82-87. 1991. 11. Jackwood, M W., Η. M Kwon, and D. A. Hilt. Infectious bronchitis virus detection in allantoic fluid using the polymerase chain reaction and a DNA probe. Avian Dis. 36:403-409. 1992. 12. Jackwood, M W., N. Μ H. Yousef, and D. A. Hilt. Further development and use of a molecular serotype identification test for infectious bronchitis virus. Avian Dis. 41:105-110. 1997. 13. Karaca, K., S. Naqi, and J. Gelb, Jr. Production and characterization of monoclonal antibodies to three infectious bronchitis virus serotypes. Avian Dis. 36:903-915. 1992.

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33 TURKEY CORONAVIRUS Mark W. Jackwood and James S. Guy

SUMMARY. Turkey coronavirus (TCoV) is the cause of coronaviral enteritis in turkeys. TCoV produces an economically significant enteric disease in commercial turkeys, and has been associated with poult enteritis and mortality syndrome (PEMS). Young turkeys are affected and diarrhea is the most common clinical sign. Agent Identification. Virus can be isolated from fecal droppings, cloacal swabs, intestinal contents, intestines and the bursa of Fabricius. The virus is propagated in embryonating eggs and identified by electron microscopy, fluorescent antibody tests, immunohistochemistry, antigen capture enzyme linked immunosorbent assay (ELISA), or reverse transcriptase-polymerase chain reaction (RT-PCR). Serologic Detection in the Host Antibodies against TCoV can be detected by the indirect fluorescent antibody test, commercial ELISA for infectious bronchitis virus (IBV) or competitive ELISA. TCoV is closely related to, and should be differentiated from IBV as well as a number of other viruses that cause enteric disease in turkeys. affected. Characteristic microscopic lesions of the intestinal tract are an increase in crypt depth, a decrease in villous length, a decrease in intestinal diameter, cuboidal epithelium with a loss of microvili, and an increase in heterophils and lymphocytes in the lamina propria (13). In the bursa of Fabricius, characteristic microscopic lesions consist of epithelial necrosis with replacement of the normal pseudostratified columnar epithelium with a stratified squamous epithelium (13).

INTRODUCTION Turkey poult enteritis is a severe and extremely important disease in commercial turkeys because it can lead to enormous economic losses. An enveloped positive-stranded RNA virus designated turkey coronavirus (TCoV) is the etiologic agent of turkey coronaviral enteritis. The original TCoV described in Minnesota in 1951 (TCoV [Minnesota]), which causes bluecomb disease, initially was believed to be similar to bovine coronavirus, a group Π coronavirus (9, 22, 24). However, recent studies have shown that TCoV (Minnesota) and other TCoV isolates associated with enteritis and PEMS in turkeys are antigenically and genomically similar to each other and to a group ΙΠ coronavirus, infectious bronchitis virus (IBV) (3,10, 11). In 1997, Guy et al. (10) reported that a TCoV isolate from North Carolina turkeys (NC95) was similar to other TCoV isolates as well as TCoV (Minnesota), and these viruses likewise were similar to IBV. Their conclusions were based on cross reactivity of polyclonal antibodies in the fluorescent antibody test and immunoperoxidase procedures. In addition, they found that a monoclonal antibody directed against the membrane protein of IBV reacted with the TCoV. Since that report, several other studies conducted on TCoV isolates, showed that the order of the genes at the 3’ end of the genome was similar to IBV (1, 3, 15, 16, 21) reinforcing the fact that IBV and TCoV are closely related. The most important structural protein in coronaviruses is the spike glycoprotein. The S glycoprotein forms club shaped projections on the surface of the virus particles. It is anchored in the envelope of the virus and consists of two subunits designated SI and S2. Spike mediates virus attachment to the host cell and is for the most part responsible for host cell specificity. In addition neutralizing antibodies are directed against the SI subunit of spike (2). Published sequence data for the TCoV spike gene (GenBank Acc. Nos. AY342356 and AY342357) shows that TCoV spike and IBV spike are clearly different (less than 23% amino acid similarity). The TCoV spike sequence was also important in establishing that different isolates of TCoV appear to be genetically similar (greater than 90% similarity).

SAMPLE COLLECTION

Samples for TCoV isolation include fecal droppings, cloacal swabs, intestinal contents, intestines, or bursa of Fabricius. Because coronaviruses are extremely labile, only fresh droppings should be collected and the samples should be kept cold (on ice at 4 C or frozen) at all times. Cold phosphate buffered saline (PBS) pH 7.4 or minimal essential medium should be added to the clinical samples and mixed well or homogenized if the samples are tissues. Next the samples are clarified by centrifugation, and filtered through a 0.45 pm filter and stored frozen, preferably at -80 C. PREFERRED CULTURE MEDIA AND SUBSTRATES

The preferred laboratoiy host system is specific-pathogen-free (SPF) embryonating turkey eggs between 15 and 21 days of incubation. Commercial eggs can be used as long as they do not contain TCoV specific antibodies. Chicken embryonating eggs between 16 and 18 days of age can also be used to propagate the virus. Eggs are inoculated via the allantoic cavity and virus is collected from the gastrointestinal tract at 48 to 72 hr post­ inoculation. Attempts to grow TCoV in cell culture have been unsuccessful. The Bluecomb virus, TCoV (Minnesota), was reported to grow in HRT-18 cells, a human rectal adenocarcinoma cell line (7); however, recent studies have failed to verify this (10). AGENT IDENTIFICATION

Electron microscopy. The virus can be identified by negative contrast electron microscopy or immune-electron microscopy. The characteristic 60 to 180nm pleomorphic, enveloped particles with club-shaped projections (spikes) can be observed in intestinal contents, intestines, and bursa of Fabricius. However, virus particles can be difficult to identify in “dirty samples” and it is recommend that the source of virus be from the embryonic gastrointestinal tract following propagation of the virus in embryonating eggs. Fluorescent antibody test and immunohistochemistry. Direct and indirect fluorescent antibody (FA) tests have been described for detection of TCoV in intestines, bursa of Fabricius, and embryoinfected gastrointestinal tissues (4, 18). Frozen tissue sections are fixed in ice cold acetone for 10 min and stored at 4 C or immediately stained with anti-TCoV antibodies directly conjugated

CLINICAL DISEASE

Young turkeys less than 4 wk of age are most susceptible to the disease and clinical signs are usually observed from 2 to 3 days post-infection (14). Clinical signs can occur suddenly and consist of watery, frothy droppings that may contain mucus and urates (13). Birds often show depression, anorexia, dehydration, and weight loss. Morbidity is near 100% and mortality can be variable depending on the age of the bird, secondary pathogens, management, and environmental conditions (13). Lesions associated with the disease are pale, thin-walled and flaccid intestines, with frothy watery contents (14). Most all of the intestinal tract can be affected but usually the lower gut, including the ceca are commonly 150

Chapter 33

with fluorescein isothiocyanate (FITC) for direct FA or for indirect FA, anti-TCoV antibodies are detected with the appropriate secondary anti-gammaglobulin conjugated with FITC. Direct and indirect FA tests have been shown to be useful for detection of viral antigens in tissues and antibodies in serum, respectively (18). The direct FA test, using FITC-conjugated turkey anti-TCoV antibodies, was found to be suitable for detection of viral antigens during the acute stages of the disease, whereas the indirect FA test was useful for detection of anti-TCV antibodies, and for monitoring flocks for exposure to TCoV (18). It was reported that TCoV can be detected from 1 to 35 days post exposure when monoclonal antibodies are used in the FA test (4). The sensitivity of that test was 69% and the specificity was 96% when compared to virus isolation. Immunohistochemical procedures (indirect FA and indirect immunoperoxidase procedures) also were used to detect the virus in frozen tissue sections prepared from intestinal tissues and bursa of Fabricius (4). Frozen tissue sections were fixed in ice-cold acetone and stored at 4 C prior to immunological staining. Indirect FA utilized TCoV-specific monoclonal antibodies and FITC-conjugated anti-mouse IgG. The indirect immunoperoxidase procedure utilized TCoV-specific monoclonal antibodies and a commercially available avidin-streptavidin immunoperoxidase kit. The virus was detected from 1 to 35 days post exposure, and the sensitivity of the test was 61%, whereas specificity was 96% when compared to virus isolation. Antigen capture enzyme linked immunosorbent assay. Using TCoV-specific antisera prepared in rabbits and guinea pigs, a double-antibody ELISA was developed to detect TCoV in intestinal contents (8). The test was found to be more sensitive than electron microscopy for detection of TCoV. Additionally, the test was not specific for TCoV as IBV was also detected. Briefly, TCoV-specific antibody is coated onto a 96 well ELISA plate and intestinal contents from turkey poults with diarrhea are added to the well. After incubation, the plate is washed with PBS and secondary antibody specific for TCoV conjugated to peroxidase is applied. Following incubation, the plate is again washed and the appropriate substrate applied to detect the virus. Reverse transcriptase-polymerase chain reaction. Several RTPCR tests have been developed to detect TCoV in intestinal contents, fecal droppings, and tissues. All of the tests reported to date cross-react with IBV. Breslin et al. (4) reported an RT-PCR test for TCoV that amplifies a 1100 bp region spanning the matrix and nucleocapsid genes. The sensitivity of the test was 160 ΕΠ)50 and the identity of the amplified product was confirmed by nucleic acid sequencing. Velayudhan et al. (23) used 3 sets of primers designed to amplify the polymerase gene and the nucleocapsid gene of TCoV. The sensitivity was between 1 and 10 ng/ml of RNA and cross-reactions with IBV were identified with another set of primers specific for the spike gene of IBV. Finally Sellers et al. (19) developed a multiplex RT-PCR test that amplifies the nucleocapsid gene of TCoV as well as IBV, and Spackman et al. (20) developed a real-time RT-PCR test that amplifies the matrix protein gene of both TCoV and IBV. Sensitivity of the multiplex and real-time tests was from 1-5 ng of RNA and 1100 gene copy numbers respectively. Generally, the RT-PCR test can detect TCoV between 1 and 14 days post exposure in cloacal swabs and intestinal tissues with cloacal swabs providing more consistent results. Procedures for extraction of viral RNA vary but the most widely used is Trizol LS reagent (Invitrogen, Inc., Carlsbad, CA); this reagent is used according to the manufacturer’s instructions. The RT-PCR reaction conditions vary with each set of primers but in general follow the recommendations of the RT-PCR kit manufacturers and consist of a 30 to 60 min reverse transcription step followed by heat denaturation for 5 min and 35 to 40 PCR cycles. The PCR products are observed on an agarose gel following electrophoresis or are sequenced. The real-time RT-PCR products were detected with a specific probe labeled with a fluorescent dye.

Turkey Coronavirus

SEROLOGIC DETECTION IN THE HOST

The indirect fluorescent antibody test (IFAT) has been established to detect antibodies to TCoV (5, 18). Antigen for this test consists either of frozen sections of TCoV-infected embryo intestines or epithelium exfoliated from bursa of Fabricius of infected turkeys (13). Frozen tissue sections are prepared from intestinal tissues of TCoV-infected turkey embiyos, 24-48 hr after inoculation with embryo-adapted TCoV strains. Bursa of Fabricius epithelial cells from experimentally infected turkeys are spotted onto microscope slides, air-dried, and fixed in ice-cold acetone. Dilutions of sera are prepared in PBS and applied to the slides. Following incubation, the slides are washed in PBS and incubated with FITC conjugated anti­ turkey IgG (H&L, Kirkegaard & Perry Laboratories, Gaithersburg, MD). The slides are washed and examined for fluorescence with a microscope containing an ultraviolet light source. Serodiagnosis of TCoV can be demonstrated by ascending serum antibody titers using the ELISA test. A commercially available ELISA test for IBV (IDEXX, Westbrook, Maine) cross-reacts and can be used to detect TCoV serum antibodies in turkeys when a conjugated secondary antibody against turkey immunoglobulin (goat anti-turkey IgG H&L, Kirkegaard & Perry Laboratories, Gaithersburg, MD) is used (17). A competitive ELISA test was also reported. That test utilizes a recombinant baculovirus expressed TCoV nucleocapsid protein and a biotin-labeled monoclonal antibody against the TCoV nucleocapsid protein (12). The ELISA tests are reported to detect TCoV antibodies beginning on day 10 through day 28 post exposure in experimentally infected turkeys (12), and both tests cross-react with IBV antibodies. DIFFERENTIATION FROM CLOSELY RELATED AGENTS Differentiation of TCoV from IBV infection is important because of the close relationship between those two viruses. TCoV has been detected for up to 14 days post exposure in the gastrointestinal tract of experimentally infected chickens by RT-PCR (14). IBV, however, has a strict host range infecting only chickens and pheasants (6). In addition, TCoV has a tropism for intestinal and bursa of Fabricius epithelium of turkeys, whereas IBV typically infects the upper respiratory tract, kidneys and reproductive tract of chickens (11). Other causes of turkey enteric disease include a number of viruses including turkey astrovirus, reovirus, rotavirus, and enterovirus, as well as bacteria, protozoa and fungi. Differentiation of TCoV from those infectious agents can be accomplished using the tests described above. REFERENCES

1. Akin, A., T. L. Lin, C. C. Wu, T. A. Bryan, T. Hooper and D. Schrader. Nucleocapsid protein gene sequence analysis reveals close genomic relationship between turkey coronavirus and avian infectious bronchitis virus. Acta Virol. 45:31-38. 2001. 2. Boursnell, Μ, Μ M Binns, T. Brown, D. Cavanagh and F. M Tomley. Molecular biology of avian infectious bronchitis virus. Karger,New York 0:65-82. 1989. 3. Breslin, J. J., L. G. Smith, F. J. Fuller and J. S. Guy. Sequence analysis of the turkey coronavirus nucleocapsid protein gene and 3' untranslated region identifies the virus as a close relative of infectious bronchitis virus. Virus Res. 65:187-193. 1999. 4. Breslin, J. J., L. G. Smith, H. J. Bames and J. S. Guy. Comparison of virus isolation, immunohistochemistry, and reverse transcriptase-polymerase chain reaction procedures for detection of turkey coronavirus. Avian Dis. 44:624-631. 2000. 5. Breslin, J. J., L. G. Smith and J. S. Guy. Baculovirus expression of turkey coronavirus nucleocapsid protein. Avian Dis. 45:136-143. 2001. 6. Cavanagh, D. and S. A. Naqi. Infectious bronchitis. In: Diseases of poultry. 11th Saif, Y. Μ, H J. Bames, J. R. Glisson, A. M Fadly, L. R. McDougald, and D. E. Swayne eds. Iowa State University Press Ames, IA. 101-119. 2003.

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17. Loa, C. C., T. L. Lin, C. C. Wu, T. A. Bryan, H. L. Thacker, T. Hooper and D. Schrader. Detection of antibody to turkey coronavirus by antibody­ capture enzyme-linked immunosorbent assay utilizing infectious bronchitis virus antigen. Avian Dis. 44:498-506. 2000. 18. Patel, B. L., E. Gonder and B. S. Pomeroy. Detection of turkey coronaviral enteritis (bluecomb) in field epiomithics, using the direct and indirect fluorescent antibody tests. Am. J. Vet. Res. 38:1407-1411. 1977. 19. Sellers, H. S., M D. Koci, E. Linnemann, L. A. Kelley and S. SchultzCherry. Development of a multiplex reverse transcription-polymerase chain reaction diagnostic test specific for turkey astrovirus and coronavirus. Avian Dis. 48:531-539. 2004. 20. Spackman, E., D. Kapczynski and H. Sellers. Multiplex real-time reverse transcription-polymerase chain reaction for the detection of three viruses associated with poult enteritis complex: Turkey astrovirus, turkey coronavirus, and turkey reovirus. Avian Dis. 49:86-91. 2005. 21. Stephensen, C. B., D. B. Casebolt and N. N. Gangopadhyay. Phylogenetic analysis of a highly conserved region of the polymerase gene from 11 coronaviruses and development of a consensus polymerase chain reaction assay. Virus Res. 60:181-189. 1999. 22. Tijssen, P., A. J. Verbeek and S. Dea. Evidence of close relatedness between turkey and bovine coronaviruses. Adv. Exp. Med. Biol. 276:457460. 1990. 23. Velayudhan, B. T., H. J. Shin, V. C. Lopes, T. Hooper, D. A. Halvorson and K. V. Nagaraja. A reverse transcriptase-polymerase chain reaction assay for the diagnosis of turkey coronavirus infection. J. Vet. Diagn. Invest. 15:592-596. 2003. 24. Verbeek, A., S. Dea and P. Tijssen. Genomic relationship between turkey and bovine enteric coronaviruses identified by hybridization with BCV or TCV specific cDNA probes. Arch Virol. 121:199-211. 1991.

7. Dea, S., S. Garzon and P. Tijssen. Isolation and trypsin-enhanced propagation of turkey enteric (bluecomb) coronaviruses in a continuous human rectal adenocarcinoma cell line. Am J. Vet. Res. 50:1310-1318. 1989. 8. Dea, S. and P. Tijssen. Detection of turkey enteric coronavirus by enzyme-linked immunosorbent assay and differentiation from other coronaviruses. Am. J. Vet. Res. 50:226-231. 1989. 9. Dea, S., A. J. Verbeek and P. Tijssen. Antigenic and genomic relationships among turkey and bovine enteric coronaviruses. J. Virol. 64:3112-3118. 1990. 10. Guy, J. S., H. J. Bames, L. G. Smith and J. Breslin. Antigenic characterization of a turkey coronavirus identified in poult enteritis- and mortality syndrome-affected turkeys. Avian Dis. 41:583-590. 1997. 11. Guy, J. S. Turkey coronavirus is more closely related to avian infectious bronchitis virus than to mammalian coronaviruses: A review. Avian Pathology 29:207-212. 2000. 12. Guy, J. S., L. G. Smith, J. J. Breslin and S. Pakpinyo. Development of a competitive enzyme-linked immunosorbent assay for detection of turkey coronavirus antibodies. Avian Dis. 46:334-341. 2002. 13. Guy, J. S. Turkey coronavirus enteritis. In: Diseases of poultry. 11th ed. Saif, Y. M, Bames, H. J., Glisson, J. R, Fadly, A. M, McDougald, Swayne, D. E. eds. Iowa State Press Ames, Iowa. 300-307. 2003. 14. Ismail, Μ M., A. Y. Tang and Y. M. Saif. Pathogenicity of turkey coronavirus in turkeys and chickens. Avian Dis. 47:515-522. 2003. 15. Lin, T. L., C. C. Loa and C. C. Wu. Existence of gene 5 indicates close genomic relationship of turkey coronavirus to infectious bronchitis virus. Acta Virol. 46:107-116. 2002. 16. Lin, T. L., C. C. Loa and C. C. Wu. Complete sequences of 3' end coding region for structural protein genes of turkey coronavirus. Virus Res. 106:61-70. 2004.

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34 ENTERIC VIRUSES Don Reynolds and Ching Ching Wu

SUMMARY. With the exception of hemorrhagic enteritis virus (covered elsewhere) enteric viruses of poultry occur primarily in young poultry. Characteristic clinical signs include diarrhea, anorexia, litter eating, ruffled feathers and poor growth. Intestines may have lesions; intestines are typically dilated and are filled with fluid and gaseous contents. The sequela to clinical disease is often stunting and / or ranting of the birds resulting in uneven flocks. Various viral agents (or combinations of these agents) can cause enteric disease and / or have been identified from the intestinal contents of birds inflicted with enteric disease. Those viruses that cause and / or are associated with enteric diseases of young poultry include astrovirases, coronaviruses, enteroviruses, rotaviruses and torovirus. Agent Identification. A number of techniques have been used to detect enteric viruses. Many of the enteric viruses were first identified by the direct visualization of the virion using electron microscopy (EM). Techniques that employ EM still remain the method of choice (or may be the only method available) for detecting many of the enteric viruses (31). Immune electron microscopy (IEM) has been advantageous for identifying many enteric viruses, especially those viruses for which physicochemical characteristics are little known. The expense, availability, and experience of EM operator have limited the use of EM for the routine diagnosis of enteric viruses. Other techniques such as fluorescent antibody (FA), genome electropherotyping, enzyme-linked immunosorbent assay (ELISA), and hemagglutination, have also been used for the detection of some of the enteric viruses. Advances in molecular biology and genomic sequencing have allowed for the reverse transcriptase polymerase chain reaction (RT-PCR) to be developed and used effectively for detecting some of the enteric viruses. Serologic Detection in the Host. Detection of antibodies is used for the diagnosis of certain enteric virus infection (VN, ELISA).

INTRODUCTION

SAMPLE COLLECTION

Enteric diseases that occur in young birds (less than 6 wk of age) are problematic for poultry producers. A number of viruses have been associated with enteric disease of young birds (35). Some of these viruses are enteropathogens. However, the role that other viruses play in enteric disease has yet to be determined. For example, in turkeys a disease once known as ’’bluecomb disease” is caused by a coronavirus. However, a similar condition in turkeys known as "poult enteritis" or "turkey viral enteritis" has been associated with other viruses including rotavirus and astrovirus infections. A similar condition in chickens that has been commonly referred to as "malabsorption syndrome," "stunting syndrome," etc., has been associated with numerous viral agents, including reoviruses and enteroviruses. Within the past decade, poult enteritis and mortality syndrome (PEMS) has been determined to be an infectious disease perhaps involving enteric viruses. Those enteric viruses that have been associated with PEMS include coronaviruses and astrovirases (4).

Intestinal samples collected from birds experiencing clinical disease (diarrhea, etc.) are of greatest benefit for isolating and identifying most enteric viruses. The intestinal tract should be removed from the bird in its entirety starting from the point where the duodenum joins the ventriculus and continuing distally to the cloaca (vent); that is, the entire intestinal tract distal to the ventriculus. The number of tracts needed depends upon the age and size of the birds involved. Typically, intestinal tracts from five to 10 birds are adequate. Ihe intestinal tracts can be placed in any container free from viral contaminants. A sterile plastic bag such as a Whirl Pac® (Nasco, Fort Atkinson, WI) is convenient. The intestinal tract should be cut into small sections of 2-5 cm in length when placing it into the sterile container (Whirl Pac®). This allows the samples to be processed easier and aids obtaining intestinal contents. The intestinal samples can then be stored and/or shipped by freezing at temperatures of -20 C or lower. Intestinal contents are an important source of virus-infected material. Although it is not necessary to ligate the intestinal tract, no attempt should be made to empty the intestinal tract of its contents. Intestinal tracts that have been fixed in formalin or other fixatives prove to be of limited value, because fixation usually results in inactivation of viruses. Once intestinal tracts are frozen, they can be stored for indefinite periods of time, as freezing appears to have minimal effects on most enteric viruses. Samples can be removed from the freezer at any convenient time for further processing. Processing the intestinal samples for further virus isolation and identification involves the following steps: 1) Thaw intestinal samples slowly, preferably at 4 C. In some instances (nonenveloped viruses) freeze-thawing multiple times aids in rupturing intestinal cells that harbor virus and breaks down tissue for easier handling of the sample. 2) Dilute the samples with sterile phosphate-buffered saline (PBS; pH 7.4) to a workable solution. Other diluents can be used; however, PBS is economical and convenient. The extent of dilution necessary depends on the sample. If the sample consists of watery diarrhea, it will take less dilution than a sample consisting of solid fecal material. Typically, a dilution of 1:5 to 1:10 (sample: diluent) is adequate. 3) Homogenize the sample. This may be accomplished in a number of ways. It is most convenient to place the plastic bags containing the sample and diluent in a stomacher for 3 min and allow the stomacher to homogenize the sample. By this method, the sample does not leave its original container (Whirl Pac®), thus minimizing the risk of contaminating the sample, laboratory, and personnel. Numerous samples may be done quickly without the

CLINICAL DISEASE

The clinical disease varies with respect to the severity of clinical signs and symptoms depending on the agent(s) and the avian species involved. However, there are a number of characteristics that are common among all the enteric virus infections. The clinical disease most commonly occurs within the second or third week of life and lasts 10 to 14 days. Therefore, most birds that contract enteric viral disease recover from the clinical disease by 6 wk of age. Characteristic clinical signs and symptoms most often include diarrhea (watery droppings), anorexia, litter eating, listlessness, and various types of enteritis. The intestinal tracts are often filled with watery contents, gas, and, in some instances, partially digested feed. The ceca are usually dilated and filled with gaseous, frothy contents. Mortality is variable and dependent upon the agent(s) and avian species involved. Generally, mortality is only slightly to moderately increased. However, with coronavirus infections of turkeys, losses of 50% or more have been reported especially in PEMS cases (4, 11). Morbidity, as a result of decreased growth (stunting), is of primary concern. Typically, 5% to 20% of a flock that has experienced enteric viral disease becomes stunted and remains so throughout the growout period.

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necessity of cleaning equipment, etc. Other methods to homogenize tissue such as using tissue grinders, blenders, or digitally expressing the tracts by hand can be as effective. Another convenient method is to “roll” the samples contained within a plastic bag with a small wall paper roller that can be obtained at a local home improvement store. 4) Sonicate samples in an ice bath for at least 1.5 min. Sonication should be accomplished by 30-sec intervals of sonication followed by 30-sec intervals of rest (no sonication) to allow heat dispersion. Sonication aids in liberating viruses from tissue and organic material. It also aids in dispersing aggregates of viruses. One must be careful when using this step, because sonication may affect membraned viruses. Additionally, sonication detaches pili from bacteria that are present in the sample. Depending on the intended use of the sample, the pili may pose potential problems. Therefore, one may elect to omit sonication or sonicate after bacteria have been removed. 5) Clarify the sample by subjecting it to centrifugation between 400 and 500 x g. This clarifies the sample of most of the

particulate matter. The supernatant is saved and used for further processing. The pellet should not be discarded, because it may retain appreciable amounts of virus. 6) The supernatant is filtered through a 0.45 pm filter. Usually, it is advisable to filter the supernatant through larger filters (such as 1.2 pm, 0.8 pm or 0.65 pm) before attempting to filter through a 0.45 pm filter. Positive pressure syringe filters are useful for this procedure. If the samples are difficult to filter, the clarification step (see step 5 above) can be repeated increasing the centrifugation force up to 5,000 x g. The sonicated filtrate can be frozen and stored. Repeated freezing and thawing may have some unwanted effect on some viruses. Therefore, it is advisable to aliquot this material in several containers before freezing. The collection of additional tissues or the use of other procedures may be of greater benefit depending upon the specific agent being sought.

Astroviruses PREFERRED CULTURE MEDIA AND SUBSTRATES

Reverse transcription-polymerase chain reaction (RT-PCR) assay A RT-PCR assay was described in 2000 by Koci et al (19, 20). PCR primers were designed to amplify two fragments, an 802 bp fragment of viral polymerase gene amplified by primers MKCap8 (TCATCATCCTCTCACACTGG) and MKCapl9 (AGCAGCAGTAGGTGGCAGTG) and 849 bp fragment of viral capsid gene amplified by primers MKPollO (TGGCGGCGAACTCCTCAACA) and MKPolll (AATAAGGTCTGCACAGGTCG). Total RNA isolated from the intestines and/or feces (experimentally infected or commercial turkeys with an acute enteric disease) using the TRIzol total RNA isolation reagent (Life Technologies, Rockville, MD) were subjected to reverse transcription to cDNA using SuperScript Reverse Transcriptase (Life Technologies). The cDNA was used as target in the PCR amplification using the above-mentioned primers. The PCR products were electrophoresised in a 1.1% agarose gel in TAE buffer, stained with ethidium bromide, and visualized by UV light. This RT-PCR assay has specificity for TAstV.

Chicken astroviruses have been reported to have been propagated in chick embryo liver cells (CEL, see Chapter 43 on primary cell culture for procedures). Briefly, CEL cells were grown to near confluency in 6mm petri dishes (5). The inocula (0.2 ml sonicated filtrate, as indicated above) can be adsorbed onto the monolayer with gentle rocking motions followed by the addition of media. Cytopathic effect (CPE) was observed after 6 days of incubation at 5% CO2 and 38.5 C. AGENT IDENTIFICATION

Astroviruses are small round, non-enveloped viruses that typically measure 28 to 30 nm in diameter (5) (with a positive-stranded RNA genome). They have characteristic five-pointed or six-pointed starlike surface projections detected by negatively stained electron microscopy (EM). The family Astrovirdidae is divided into 2 genera: mammalian and avian astroviruses. Avian astroviruses have been associated with enteric diseases in turkeys causing diarrhea and high mortality. Astroviruses were first reported in turkeys of age 6-11 days. Two types of turkey astroviruses (TAstV) have been identified TAstV-1 and TAstV-2. These two types differ immunologically and genetically from the other avian astroviruses, avian nephritis virus, duck virus hepatitis type 2 and chicken astrovirus (34). The complete genomic sequence of TAstV was reported by Koci et al (18, 20). The genome is 7,325 nucleotides and contains three ORFs; ORF la, ORF lb and ORF2, and there is a frame shift between ORF la and ORF lb. The protein products for ORF la, and ORF lb are unknown as yet, but are predicted to be non-structural proteins. It is possible that the product of ORFa is a serine protease and the product of ORF lb is a RNA-dependent RNA polymerase (5, 20). ORF2 encodes the viral capsid protein of 73-80 kDa. Classical tests to detect astroviruses include electron microscopy, fluorescent antibody detection and agar gel diffusion assay. These tests, while more user friendly, lack the required sensitivity for accurate diagnosis. Recent advances in molecular biology have opened the door for the development of many diagnostic assays for the detection of avian astroviruses and are described here.

Multiplex RT-PCR A multiplex RT-PCR was described for the detection of turkey astrovirus and turkey coronavirus TCoV (37). Intestine and feces from commercial turkey flocks and from turkey embryo intestines inoculated with TCoV orTAstV-2 were used to develop the assay. PCR primers (TCVnucleo forward, GGTAGCGGTGTTCCTGA, and TCVnucleo reverse, CCCTCCTTACCTTTAGT) for the detection of TCoV were designed to amplify a 598 bp fragment within the nucleocapsid gene of TCoV. PCR primers for TAstV-2 were designed to amplify the 802 bp fragment of viral polymerase gene as mentioned above. The amplified TCoV nucleocapsid gene sequence is conserved with nucleocapsid gene sequence of infectious bronchitis virus (IBV). This assay was specific for TAstV-2, TCoV, and IBV. The sensitivity or the detection limit of the multiplex RT-PCR was between 5 to 10 ng. In situ hybridization In situ hybridization was described in 2002 by Behling-Kelly et al (6), in which tissue samples were taken from experimentally inoculated 3-day-old turkey poults with TAstV-2. Tissues assayed were intestines, spleen, bursa and thymus. Tissues were de­ paraffinized, digested with proteinase K, and hybridized overnight with a digoxigenin-labeled riboprobe. The riboprobe was generated by ZtawHI digested plasmid p25.5, which contains a 1.5 kb segment of the extreme 3' end of the TAstV-2 genome. In vitro transcription

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Chapter 34 Enteric Viruses

with T7 RNA polymerase and digoxigenin-labeled UTP was used as an antisense riboprobe (1.6 kb in length). The in situ hybridization assay is used to detect viral replication in tissues. It was able to detect TAstV-2 in duodenum, cecal tonsils, jejunum, distal small intestine and large intestine, but not in bursa, thymus or spleen. TAstV-2 was detected after 1 day post inoculation (PI) in cecal tonsils and distal small intestine, then in all other tissues on day 3 PI. The signals were extensive in distal small intestine and cecal tonsils. The signals were relatively infrequent by day 9 PI.

day 1 to day 14 PI and the highest proportion of poults positive for virus detection was between days 4 and 9 PI.

SEROLOGICAL DETECTION IN THE HOST Neutralization test: A virus neutralization assay was described recently for Astrovirus (5). In this microtitre neutralization test, two-fold dilutions of serum were mixed with an equal volume of virus diluted in Eagle’s modified essential medium to give 200 median tissue culture infective doses per 0.1 ml, the sera were incubated at room temperature for 1 h and 0.1 ml from each dilution was placed into four microtitre wells in a 96-well plate and 0.1 ml growth medium and 3xl04 LMH cells. A virus titration series and negative serum were included in each test. The plates were incubated for 5 days at 38.58 C in a CO2 atmosphere and then read microscopically. The serum titer was the dilution where only 50% of wells showed CPE. Also, a virus neutralization test in embryonated turkey eggs was described by Tang et al (39), in which 100 EID5o of TAstV 1987 isolate and TAstV 2001 isolate were mixed with six 10-fold serial dilutions of guinea pig antisera, then inoculated into 22-day-old turkey embryos by the amniotic route. After 96 h of incubation at 37 C, embryos were examined for intestinal lesions. Enzyme-linked immunosorbent assay. An ELISA system was developed by Tang et al (39). This system is based on the purified turkey astrovirus isolates (TAstV1987 and TAstV2001). The purified antigens were used at 2g/ml in 0.05 M carbonate­ bicarbonate coating buffer and, after blocking step with 3% bovine serum albumin in phosphate-buffered saline with 0.1 Tween-20 (PBS-T), anti- TAstV 1987 and TAstV 2001 were added in 1:400 dilution, then goat anti-guinea pig IgG (H+L) horseradish peroxidase labeled antibody diluted in 1:20,000 was added, then substrate 3,3’, 5,5’-tetramethylbenzidine (TMB) mixed with equal volumes of peroxidase solution B was added for color development at 405 nm.

Multiplex real time RT-PCR A multiplex real time RT-PCR test was recently developed (38) for the detection of TAstV-2 and TCoV. Virus detection was evaluated with samples collected from poults inoculated at 1 day of age with each virus. Cloacal swabs and intestinal samples were obtained at 1,2, 3, 4, 6, 9, 14, 17, and 21 days after inoculation and also from field samples. Primers for the real time RT-PCR for astro viruses were designed to amplify a 112 bp fragment of the polymerase gene, the sequence of the forward primer is ( TAV 4248F) 5’-TCC TCC ATG ATT CTC ATA AG- ‘3 and for the reverse primer (TAV 4360 R) 5’-CTT GAC CTG GCA AAC T-‘3 and the probe sequence (TAV 4274 PB) 5’- [4J-AAG ATG CGG CGC TTG TA-{TAMRA}-‘3. Primers for TCoV were designed to amplify a 110 bp fragment of the matrix gene and their sequences are: forward primer (TCoV 2F) 5’- AGT GGC TTG CTA AGT-‘3, for the reverse primer (TCoV 112R) 5’-GCT TTG GTC ACC AGT‘3 and for the probe (TCoV 51 PB) 5’-{TXRed}-TAT GCA CAC CGG ATA GAC G-{BHQ-2}-‘3. Assay sensitivity was determined using in vitro transcribed RNA and varied by target between 150 gene copies for TAstV-2 alone and 2200 gene copies for TCoV when multiplexed. The TAstV2/TCoV test was able to detect the astrovirus isolate (NC/96) and five additional field isolates as well as TAstV-2. In the poults experimentally inoculated with TAstV-2, the virus could be detected in both cloacal swabs and intestinal tissue from

Small Enteric Viruses are mounted on N-aminoethylaminopropyltrimethoxysilane treated glass slides, processed through xylene-ethanol, and digested with 0.1% trypsin. The tissue sections are treated with methanol and 3% peroxide to quench endogenous peroxidase. Normal goat serum is used as blocking agent at a 1:10 dilution. An avidin-biotin block was used to block endogenous biotin in the tissue sections. Unlabelled goat anti-turkey IgG was added to prevent binding of the biotinylated goat anti-turkey IgG to IgG -producing cells. Hyper immune turkey anti-enterovirus serum adsorbed with intestinal powder was added in 1:160 dilution and followed by biotinylated goat anti-turkey IgG. The substrate ExtrAvidin peroxidase conjugate is added followed by another substrate 3-amino-9ethylcarbazole. Sections were counterstained with Mayer’s hematoxylin then mounted in glycerol gelatin and examined by light microscopy. Positive tissue samples were from ileum, jejunum and duodenum, with the most severely affected tissues being jejunum and ileum (16).

PREFERRED CULTURE MEDIA AND SUBSTRATES

Enteroviruslike particles from chickens can be serially propagated by inoculating the chorioallantoic membrane (CAM) of embryonated chicken eggs. Enteroviruslike particles from turkeys can be propagated by inoculating embryonated chicken eggs by the yolk sac route. Inocula can be prepared from intestinal content samples as described above (see Sample Collection). Procedures for CAM and yolk sac inoculation are described in Chapter 44 on virus propagation in embryonating eggs.

AGENT IDENTIFICATION Small enteric viruses (18-24 nm in diameter) are a group of small viruses that are similar in size and detected by EM but generally cannot be differentiated on the basis of morphology (14), Included in this category are enterovirus, parvovirus, calicivirus and astroviruses. Enteroviruslike virus was detected in young turkeys with enteritis. The virus can be propagated in embryonated turkey eggs, with buoyant density of 1.33 g/ml in CsCl, and has a single­ stranded RNA genome of approximately 7.5 kb. The standard diagnostic assay for the detection of small enteric viruses is EM in which the intestinal contents from affected turkeys are used to evaluate the presence of virus (15, 36).

Immunofluorescent antibody assay The immunofluorescent antibody assay (IFA) is used to detect virus in the infected tissues. Formalin-fixed, paraffin-embedded tissue sections were mounted on glass slides, deparaffinized, and digested with 0.1% trypsin. Slides were incubated with normal goat serum diluted 1:10 to reduce non-specific background staining. Hyperimmune turkey anti-enterovirus serum diluted 1:60 was added, followed by fluorescein-labeled goat anti-turkey IgG globulin (Kirkegaard & Perry Laboratories, Gaithersburg, MD). Stained sections were examined for infected cells using a

Immunohistochemistry. Paraffin embedded tissue is used to evaluate the presence of enterovirus-like particles. Embedded tissue

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Don Reynolds and Ching Ching Wu

fluorescence microscope (Nikon Optiphot; Nikon Inc., Garden City, N.Y.) (16).

Reverse transcription-polymerase chain reaction A turkey enteroviruslike virus was identified to be through cross immunofluorescence assay and RT-PCR (14). Turkey enteroviruslike particle RNA was amplified by RT-PCR with oligonucleotide primers specific for the polymerase gene (ORF lb) and the capsid protein gene (ORF2) of TAstV-2. RNA samples for the RT-PCR were extracted from turkey enteroviruslike virus infected turkey intestines. The RT-PCR products were of 802 bp and 849 bp , respectively. Sequence analysis of these RT-PCR products showed a high degree of similarity to those of TAstV-2 (ORFlb; 98.8% and ORF2; 96.9%).

Enzyme-Linked Immunosorbent Assay In 1993, an antigen-capture ELISA was developed for the diagnosis of enterovirus infection in turkey (15, 16). The samples were intestinal contents from naturally and experimentally infected turkeys. Hyper immune turkey anti-enterovirus serum in 1:1600 dilution in carbonate buffer was used. Five percent non fat dry milk was used as a blocking reagent. Fecal samples were sonicated and concentrated by centrifugation and diluted in minimum essential medium at 1:5 dilution. Guinea pig anti-enterovirus antibody was diluted at 1:800 in tris buffered saline and used as primary antibody for ELISA. Sheep anti guinea pig IgG peroxidase -labeled antibody was used at 1:1000 in tris-buffered saline as secondary antibody. 2, 2’-Azino-bis (3-ethyl-benz-thiazoline-6-sulfonic acid) in 0.05 M citrate buffer with 3% hydrogen peroxide was used as a substrate. Color development was read at 410 nm. The sensitivity and specificity for this ELISA system compared to EM were 0.963 and 879, respectively. 0.

SEROLOGICAL DETECTION IN THE HOST

Antibodies to enterovirus have been detected by serum neutralization and indirect immunofluorescence tests. Serology is useful to determine the status of enterovirus infection in specific pathogen free birds. However, due to the lack of virus isolates and reference antisera, routine serological diagnosis is not recommended (30).

Coronaviruses PREFERRED CULTURE MEDIA AND SUBSTRATES

AGENT IDENTIFICATION

There is currently no known cell culture system that is able to support turkey coronavirus (TCoV) infection and this is a significant limitation both for the ease of diagnosis and the investigation of the virus biology. Turkey coronaviruses can be propagated/isolated in embryonating chicken or turkey eggs more than 15 days of age by amniotic route of inoculation. Twenty two or 23-day turkey embryos are commonly used for isolation and propagation of TCoV (21). The inoculum is prepared from the intestines, intestinal contents, or bursal tissues of suspected turkeys or turkey poults. The intestines, intestinal contents, or bursal tissues are homogenized with 5 volumes of PBS or TBS by motar and pestle or a tissue homogenizer. The homogenates are centrifuged at 500 x g for 30 min. The supernatant is filtered through a 0.45 pm membrane filter membrane prior to inoculation into eggs. If necessary, antibiotics (penicillin or streptomycin) can be added to the filtrate (inoculum) before inoculation. In addition, the inoculum, if needed, can be activated by trypsin by adding 10 pg/ml of type ΧΙΠ trypsin to the inoculum, followed by incubation at 37 C for at least 1 hr. The air cell end of the egg is disinfected by 70% alcohol and iodine. A 26-gauge needle with the syringe containing the inoculum is inserted through a hole in the egg shell (created in the air cell end of the egg) and subsequently through the chorioallantoic membrane and reaches the amniotic cavity. One-hundred to 500 pl of inoculum are injected into the amniotic cavity. The needle is removed and the hole in the egg shell is sealed by parafilm, cellophane tape, or silicone gel. Viruses can be recovered from turkey embiyo intestines or bursae 2 to 5 days after amniotic inoculation. Turkey coronavirus is detected in turkey embryo intestines or bursae by immunofluorescent antibody assay (IFA) using a TCoV-specific antiserum or monoclonal antibody. Electron microscopy (EM) can also be used for intestinal contents or homogenates to detect enveloped viruses of 50-200 nm with typical coronavirus peplomers on the membrane. Definitive diagnosis by EM can be achieved by immune electron EM (IEM) busing a TCoV-specific antiserum.

Coronaviruses have been reported to cause clinical diseases in chickens, turkeys, ducks, geese, pheasants, and pigeons and have been isolated from these animal hosts. Unlike coronavirus induced infectious bronchitis in chickens, the primary target tissue of coronavirus in the other avian species is intestinal tract. Based on antigenic relationship between turkey coronavirus (TCoV) and other coronaviruses, TCoV was classified as a group 3 coronavirus with avian infectious bronchitis virus (IBV). The viral particles of TCoV are surrounded by a fringe of regularly spaced petal-shaped projections attached to the particles by a short stalk. Turkey coronaviruses are hemagglutinated with rabbit erythrocytes. Turkey coronavirus has the buoyant densities of 1.14 to 1.15 and 1.18 to 1.20 g/ml, respectively, using sucrose density gradient ultracentrifugation (21). Coronavirus genome contains a single, positive-strand RNA molecule, which is about 27 to 33 kilobases (KB) and has a methylated cap at the 5’ end and poly (A) tail at the 3’ end. Genome organization of coronavirus is 5’-polymerase gene-spike gene (S)membrane protein gene (M)-nucleocapsid gene (N)-3’, in which S, M, and N are the structure genes. The whole TCoV genome consists of 27,749 nucleotides, excluding poly (A) tail. Turkey coronavirus polyprotein gene encodes two open reading frames. There are 9 open reading frames in the structure protein genes, representing the entire S protein gene, tricistronic gene 3, M protein gene, bicistronic gene 5, and N protein gene in the order of 5’ to 3’(23). Turkey coronavirus can be identified and/or confirmed by EM, IEM, IFA, immunohistochemistry, hemagglutination, RT-PCR, and multiplex PCR.

Electron microscopy Intestinal contents or homogenates (3 or 5 grams) are placed into a stomacher bag with addition of 50 ml of water and the stomacher bag is processed in the stomach machine for 30 sec. Four ml of the mixture is added with an equal volume of water and centrifuged at 8,000 rpm for 5 min. Four ml of the supernatant is added with an equal volume of water and centrifuged at 20,000 rpm for 60 min. The supernatant is poured off and the pellet is added with a mixture consisting of 4 drops of phosphotungstic acid and 3 drops of 0.1 % 156

Chapter 34

bovine serum albumin and vortexed for 30 sec to 1 min. The stained pellet is transferred to a nebulizer, sprayed onto the grids, and observed on an electron microscope. Turkey coronavirus particles are spherical, enveloped, and surrounded by regularly spaced petalor pear-shaped projections. They are 50 to 200 nm in diameter (21).

Enteric Viruses

cold ethanol and pellets were resuspended in RNAse-free water (7). In another report (45), the intestines were homogenized in minimom essential medium (MEM) with penicillin (0.5 U/ml) and streptomycin (0.5 mg/ml). The homogenate (20% w/v) was centrifuged at 8,000 x g for 10 min. Viral RNA was extracted from the supernatant of intestinal homogenate using QIAamp viral RNA mini kit (Qiagen, Valencia, CA) (45). RNA was subjected to reverse transcription followed by PCR amplification. Several sets of PCR oligonucleotide primers designed from the nucleocapsid (N) protein gene sequence of TCoV were used in PCR amplification: Nl-upper (CAGCGCCAGTCATCAAAC) and N2-lower (TGGTCAAACTTGTCAGGGTCC) amplifying 380 bp (45), N3upper (CAAGTAAAGGCGGAAGAAAAC) and N4-lower (GCCTTAGTAATGCGAGAGCCC) amplifying 417 bp (45), and NF (TCTTTTGCCATGGCAAGC) and NR (TTGGGTACCTAAAAGTTCATTCTC) amplifying 1230 bp (26). However, all these primer sets also amplified N gene of infectious bronchitis virus (IBV).

Immune electron microscopy Intestines or intestinal contents are mixed or homogenized with 5 volumes of TBS and centrifuged at 250 x g for 10 min. The supernatant is filtered by 0.8-pm membrane filter, followed by 0.45-pm membrane filter. The filtrate is incubated with diluted turkey anti-TCoV serum overnight at 4 C. The samples are centrifuged at 20,000 rpm for 60 min. The pellet is resuspended in distilled water and added with a 3.0% phosphotungstic acid solution to yield a final concentration of 1.5% phosphotungstic acid. The samples are applied to the grids and observed on an electron microscope (16). Immunofluorescent antibody assay The IFA assay for TCoV antigen detection in the intestine or bursa of Fabricius is a sensitive assay due to the use of a fluorochrome conjugated antibody (14, 22, 25, 32). TCoV antigen can be detected in the intestine of experimentally infected turkeys from 1 to 28 days post infection. The small intestine (jejunum and ileum) is frozen immediately after collection, embedded in embedding medium, and sectioned in 6-pm thickness. Tissue sections are fixed in acetone for 30 min at room temperature and incubated with turkey antiserum specific for TCoV at a dilution of 1:40 in dilution buffer, containing 150 mM phosphate buffer, 0.85 % NaCl, 1 % BSA, and 0.02 % Tween-20, in a humidifying chamber at room temperature for 30 min. After washing with PBS buffer for 3 times, intestinal sections are incubated with fluorescein isothiocyanate (FITC) conjugated goat anti-turkey IgG (H+L) antibody (Kirkegaard & Perry Laboratories) at a dilution of 1:40 in dilution buffer at room temperature for 30 min. Sections are read on a fluorescent microscope (Nikon Optiphot; Nikon Inc.).

Multiplex PCR A rapid, sensitive, and specific multiplex PCR method has been established for specifically differential detection of TCoV, IBV, and bovine coronavirus (BCoV) (27). Intestines of turkey embryos infected with TCoV were homogenized with 5-fold volume of phosphate buffered saline (PBS) solution. These homogenates were centrifuged at 1,500 x g for 10 min. Two hundred microliters of virus-containing supernatants were mixed with 1 ml of RNApure reagent (GenHunter, Nashville, TN) and incubated on ice for 10 min. After addition of 180 μΐ of chloroform, the mixture was mixed vigorously for 10 sec and centrifuged at 13,000 x g for 10 min. The upper aqueous phase was mixed with equal volume of cold isopropanol and incubated on ice for 10 min. The RNA precipitate was pelleted by centrifugation at 13,000 x g for 10 min and washed with 70 % ethanol. The RNA was dissolved in 50 μΐ of diethyl­ pyrocarbonate (DEPC) treated sterile double-distilled water. The RNA was combined with random hexamers (50 ng) in 11 μΐ of DEPC-treated water, heat denatured at 70 C for 3 min, and immediately placed on ice for 5 min. Reverse transcription buffer containing 200 units of reverse transcriptase (Superscript Π system, Life Technologies, Gaithersburg, MD) and 0.2 mM of each of the four deoxynucleotide triphosphates (Promega Corp., Madison, WI) was added. A total volume of 20 μΐ reverse transcription was carried out at 42 C for 60 min. The primers used in multiplex PCR were based on the alignments of spike (S) or N gene sequences among TCoV, IBV, and BCoV to identify the variable and conserved regions (23). The upstream primer N103F and downstream primer N102R common to both TCoV and IBV were designed according to conserved regions of N gene sequences. The sequence of upstream primer N103F (cctgatggtaatttccgttggg) and that of downstream primer N102R (acgcccatccttaataccttcctc) amplify a 357-bp sequence of TCoV or IBV N gene in the conserved regions corresponding to nucleotide position 445 to 801 of TCoV N gene. The upstream primer S306F (tgtatctaatttgggtgggtttga) and downstream primer S306R (ataagctgctaattgaagggatgc) are specific to TCoV and based on the alignments of S gene sequences in the variable regions among these viruses. This set of primers specifies a 727-bp sequence corresponding to nucleotide position 2,019 to 2,745 of TCoV S gene (23). The upstream primer S3 (atgtgtgtaggtaatggtcctgg) and downstream primer S6 (agcaactacgaatcataaaa) are specific to BCoV and designed according to variable regions of S gene sequences among these viruses. This set of primers amplifies a 568-bp sequence corresponding to nucleotide position 1,488 to 2,055 of BCoV-Quebec S gene. PCR was performed in a 96-well thermal cycler (GeneAmp, Perkin-Elmer Cetus Corp., Norwalk, CT). The reaction mixtures contained 2μ1 of cDNA, 0.2 μΜ of each of the primers N103F, N102R, S306F, S306R, S3, and S6 in 50 μΐ of PCR

Immunohistochemistry Frozen intestinal or bursal sections can be acetone-fixed, incubated with TCoV-specific monoclonal antibody (Mab 4.24), and followed by an immunoperoxidae procedure to reveal TCoV antigens in the enterocytes in tissue sections on a light microscope (7). This assay had high specificity (96%) and low sensitivity (61%) when compared to virus isolation (7).

Hemagglutination Turkey coronaviruses can agglutinate guinea pig and rabbit erythrocytes. Turkey coronavirus antigens are propagated, purified, and concentrated as described in the section of virus neutralization for TCoV in this Chapter (9, 21). Erythrocyte suspension of 0.5% are used to mix with equal volume of TCoV antigen. The assay is performed at room temperature for 1 hr. The procedures of hemagglutination and hemagglutination inhibition can be referred to the Chapter on serological procedures.

Reverse transcription-polymerase chain reaction Several approaches have been used to extract viral RNA from the intestinal contents/dropping samples. In one method (7), the sample was prepared as 20% (w/v) suspensions in TNE buffer (0.01 M Tris-hydrochloride, pH7.4, 0.1 M NaCl, 1 mM ethylenediaminetetraacetic acid) and sonicated for 30 sec. These suspensions were clarified by centrifugation at 1000 x g for 10 min at 4 C and then at 8000 x g for 30 min at 4 C. The supernatant was layered onto a 20% (w/v) sucrose cushion and centrifuged at 80,000 x g for 2 hr at 4 C. The resultant pellets were incubated in 0.5% sodium dodecyl sulfate for 5 min at room temperature followed by two phenol-chloroform extractions. Nucleic acid was precipitated in 157

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plasmid vector (Invitrogen) and transformed into E. coli strain TOPI OF' (Invitrogen). Correct clones were selected and subjected to sequencing reaction using dideoxy-cycle sequencing method with the corresponding sequencing primers for both strands. The nucleotide and deduced amino acid sequence similarities among the TCoV isolates were analyzed by the Clustal W method in MegAlign module of the DNAstar program (Lasergene Corp, Madison, WI). Percent similarities were calculated to find nucleic acid and amino acid pair distances. Based on the obtained sequences, phylogenetic trees of TCoV isolates were constructed. Pair-wise comparison of nucleotide sequence distance for the entire 3’-end structural protein gene sequences among TCoV isolates revealed the similarity scores ranging from 92.7 % to 99.4 %. Pair-wise comparison of nucleotide and deduced amino acid sequence distance of S protein gene sequences among TCoV isolates had the similarity scores ranging from 93.0 % to 99.7 % at the nucleotide level or from 92.5 % to 99.3 % at the amino acid level. Phylogenetic analysis according to the entire 3’-end structural protein region or the deduced amino acid sequence of S protein gene showed that TCoV isolates were clustered within the same genomic lineage. This indicates that TCoV isolates from various geographic locations in the U.S. are closely related genetically (28).

buffer comprised of 0.2 mM of each of the four deoxynucleotide triphosphates, 5 units of Taq DNA polymerase (Promega), 10 mM Tris-HCl (pH 9.0), 50 mM KC1, 1.5 mM MgCl2, 0.1 % Triton X100, and 0.01 % gelatin. The cyclic parameters of the PCR were 94 C for 30 sec for denaturation, 50 C for 1 min for annealing, 72 C for 1 min for extension for 25 cycles followed by 72 C for 10 min final extension. Two PCR bands of 727 and 357 bp, respectively, were seen in the TCoV isolates. One PCR band of 357 bp was shown in the IBV strains. One PCR band of 568 bp was obtained for the BCoV strains and isolates.

Strain Variability Turkey coronavirus was identified in the early 1970s as the major cause of the most costly disease of turkey encountered in Minnesota between 1950s and 1970s. In addition, outbreaks of turkey coronaviral enteritis occurred in Quebec, Canada in late 1980s, Indiana, U.S. in early and mid 1990s, and North Carolina, U.S. and Virginia, U.S. in mid and late 1990s and has continued to remain as the disease of threat to the turkey industry in North Carolina, Virginia, Arkansas, Missouri, and other states in the U.S. in the 2000s. Variability of TCoV strains or isolates from different geographical locations in different time period can be determined by immunological methods and molecular approaches. Immunological methods. Immunofluorescent antibody assay, ELISA, and virus neutralization are immunological methods that can be used to study variability of TCoV strains or isolates. The procedures of these methods have been described in this Chapter. The cross antigenic reactivity of 18 TCoV isolates from various geographical locations in the U.S. was studied by IFA using antibodies to different TCoV isolates (22). Intestinal sections were prepared from turkey embryos infected with different TCoV isolates and reacted with polyclonal or monoclonal antibodies specific to TCoV in immunofluorescent antibody staining. All 18 TCoV isolates had the same antigenic reactivity pattern with the same panel of antibodies. This indicates that TCoV isolates from various geographic locations in the U.S. are antigenically identical or closely related (22). Molecular methods. Turkey coronavirus genome is a positive single-stranded capped RNA with a polyadenylated 3’ end. The 5’ two-thirds of the coronavirus genome consist of two overlapping open reading frames (ORFs) that encode non-structural proteins including the viral RNA-dependent RNA polymerase. Another onethird nucleotide sequences, including the structural protein genes and 3’ UTR, in the 3’ end have a total of 6,963 nucleotides in the structural protein genes that consist of the entire S protein gene, tricistronic gene 3, membrane (M) protein gene, bicistronic gene 5, and N protein gene in the order of 5’ to 3’ along the genome (23). PCR, cloning, sequencing, and sequence analysis (including sequence alignment, degree of sequence homology, and phylogenetic analysis) are the major molecular methods to determine variability of TCoV strains or isolates. The 3’-end structural protein gene sequences of TCoV isolates associated with outbreaks of acute enteritis in Indiana, North Carolina, and Minnesota have been studied by carrying out PCR, cloning, and sequencing (28). TCoV isolates were propagated in 22-day-old turkey embryos. Total RNA was extracted from the infected intestines and intestinal content by using guanidinium thiocyanate and acid-phenol. The RNA was dissolved in 150 μΐ of diethyl­ pyrocarbonate (DEPC) treated sterile double-distilled water. Two micrograms of the total RNA was heat denatured at 100 C for 3 min and slowly cooled to 22 C in 15 min in reverse transcription (RT) buffer (Life Technologies) containing 40 ng of random hexamers. The reverse transcription was carried out at 42 C for 60 min. Three microliters of cDNA were used in PCR amplifications with the primers spanning for 4 overlapping fragments covering the entire 3’end structural protein genes as described in a previous report (23). One microliter of the amplicon was used to ligate with pCR-Π

SEROLOGICAL DETECTION IN THE HOST Immunofluorescent antibody assay for antibody to TCoV Frozen turkey intestines infected with TCoV were sectioned at 6pm thickness and incubated with 2-fold serially diluted turkey serum at room temperature for 30 min. The intestinal sections were subsequently incubated with FITC-conjugated goat anti-turkey IgG (H+L) antibody (Kirkegaard & Perry Laboratories) at a dilution of 1:40 in dilution buffer at room temperature for 30 min in a humidifying chamber. Sections were read on a fluorescent microscope (Nikon Optiphot; Nikon Inc.). The titer of turkey sera was defined as the reciprocal of the highest dilution of test sample still having positive staining (25, 32). Alternatively, TCoV-infected epithelial cells exfoliated from the bursae of Fabricius can be served as the source of TCoV antigen. Exfoliated TCoV-infected epithelial cells were spotted onto glass slides, air-dried, and fixed in cold absolute acetone for 10 min. FITC-conjugated rabbit anti-chicken immunoglobulin G (ICN Biomedicals, Inc., Costa Mesa, CA) with a 1:40 dilution was used as the secondary antibody (13). Antibody-capture enzyme-linked immunosorbent assay Based on the positive antigenic cross-reactivity between TCoV and IBV, commercially available IBV-coated ELISA plates (IDEXX, Westbrook, ME) were successfully used for the detection of anti-TCoV antibody (24). Anti-TCoV hyperimmune turkey serum and normal turkey serum were used as positive or negative control sera for optimization of the ELISA. Goat anti-turkey IgG (H+L) conjugated with horseradish peroxidase (Kirkegaard & Perry Laboratories) was used as detector antibody (conjugate). The differentiation between anti-TCV hyperimmune serum and normal turkey serum were achieved when using 1:40 serum dilution, and 1:1,600 conjugate dilution. Ninety six-well microtiter plates coated with IBV antigens were incubated with turkey serum samples (1:40 dilution) to be tested in quadruplicate and incubated at 37 C for 1 hr. The plates were subsequently incubated with goat anti-turkey IgG (H+L) conjugated with horseradish peroxidase (1:1,600 dilution) at 37 C for 1 hr., followed by the addition tetramethyl benzidine (TMB) solution. Two reference wells that contained all reagents except serum samples were included in each plate. Positive and negative control sera as well as test sera were tested in duplicate. The absorbance value of each well was measured at 450 nm using a spectrophotometer (Vmax™ Kinetic Microplate Reader, Molecular Devices Corporation, Menlo Park, CA). The ELISA value or S/P ratio of each test serum was calculated as (absorbance

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value of sample serum minus absorbance value of negative control serum) divided by (absorbance value of positive control serum minus absorbance value of negative control serum). The optimum cutoff point to separate positivity and negativity was determined to be 0.18 by logistic regression analysis. The sensitivity and specificity of this ELISA relative to IFA test were 93.1 % and 96.7 %, respectively. Another ELISA to detect TCoV antibody (26) was developed using the recombinant N protein of TCoV expressed in E. coli transformed with pTriEx-1 (Novagen, Madison, WI) plasmid construct carrying N gene of TCoV. The optimum conditions for differentiation between anti-TCoV serum and normal turkey serum were: coating concentration of recombinant E. coli-expressed TCoV N protein at 20 pg/ml, serum dilution of testing turkey serum sample at 1:200 to 1:800, and conjugate dilution at 1:10,000 or 1:20,000. The optimum cutoff point to separate positivity and negativity was determined to be 0.18 by logistic regression analysis. In addition, Recombinant N protein of TCoV produced by recombinat baculovirus encoding TCoV N gene (rBTCV/N) has been shown to be useful in detecting antibody to TCoV in an antibody-capture ELISA (8). Recombinant baculovirus was generated by co-transfection of linearized AcMNPV DNA (Bac-NBluea DNA, Invitrogen, San Diego, CA) and pBacTCV DNA carrying TCoV N gene cloned into pMelBac C vector (Invitrogen). The rBTCV/N protein was diluted 1/80 in 0.2 M carbonate/0.2 M bicarbonate buffer, pH 9.6, added to 96-well ELISA plates, and incubated overnight at 4 C. rBTCV/N protein coated plates were washed three times with PBS plus 0.05% Tween 20 (PBST), followed by 200 μΐ of block buffer (PBST containing 1% nonfat dried milk) being added to each well and incubated for 1 hr at room temperature. Following washing three times with PBST, 75 μΐ of testing serum, diluted 1:40 in block buffer, was added to each well and incubated for 30 min at 37 C. Seventy-five μΐ of peroxidaselabeled goat anti-chicken immunoglobulin G in 1:50 dilution was added to each well and incubated for 30 min at 37 C. One-hundred μΐ of 2, 2'-azino-bis (3-ethylbenzthiazoline-6-sulfonic acid) substrate (Kirkegaard and Perry Laboratories) was added to each well; color development was stopped after 15 min with 1% (w/v) SDS in water. Plates were read with a ELISA reader at 405 nm. The TCoV N ELISA detected antibodies in all antisera prepared against TCV and IBV strains. Samples were considered positive if their absorbance was > 0.1. Absorbance values for TCoV and IBV antisera were > 0.3. Since TCoV and IBV N gene sequences are conserved and share cross-reactivity, antibody-capture ELISA based on IBV antigen or recombinant TCoV N protein can detect serum antibody to TCoV as well as that to IBV. However, S gene sequences between TCoV and IBV are markedly different, with only 33.8% to 33.9% of sequence homology (23). Therefore, antibody-capture ELISA based on TCoV S protein should be able to detect TCoV-specific antibody, differentiate between antibodies to TCoV and IBV, and evaluate TCoV-specific antibody response in the turkey flocks. A TCoVspecifc antibody-capture ELISA has been achieved for the detection of antibody to TCoV in turkey sera using recombinant partial S protein of TCoV expressed in E. coli transformed with pTriEx-1 (Novagen) plasmid construct containing the insert of partial S gene of TCoV as the coating antigen in the ELISA plates (46). A partial SI gene fragment of TCoV was ampilified by PCR, cloned into an expression vector (pTriExl, Novagen), and expressed in E. coli (Rosetta cells, Novagen). The recombinant partial SI protein was purified by His-Bind column chromatography (Novagen) and used to coat the ELISA plates. The optimal parameters for differentiation between anti-TCoV serum and normal turkey serum by ELISA were: purified recombinant partial S protein at 20 pg/ml, serum dilution 1:200 to 1:800, and conjugate dilution 1:10,000, 1:20,000, or 1:40,000. The optimum cutoff point to separate positivity and negativity was determined to be 0.32 by logistic regression method.

The sensitivity and specificity of ELISA relative to IFA test were 98.7% and 98.8%, respectively. The recombinant TCoV S protein­ based antibody-capture ELISA is useful in detection, differentiation, and evaluation of antibody response to TCoV. Competitive enzyme-linked immunosorbent assay A competitive enzyme-linked immunosorbent assay (cELISA) was developed for detection of antibodies to TCoV by using a recombinant baculovirus-expressed TCoV N protein (as mentioned above) and biotin-labeled TCoV N protein-specific monoclonal antibody (Mab 4.23) (13). The rBTCV/N protein was diluted 1:1280 in 0.2 M carbonate/0.2 M bicarbonate buffer and 75 μΐ was added to each well in 96-well ELISA plates and incubated overnight at 4 C. rBTCV/N protein-coated plates were washed three times with 0.01 M PBS, pH 7.2, containing 0.05% Tween 20 (PBST), 200 μΐ of block buffer (PBST containing 1% nonfat dry milk) was added to each well and incubated for 1 hr at 25 C. After washing three times with PBST, positive and negative control sera and test sera were diluted 1:10 in block buffer and 50 μΐ of each were placed in duplicate wells and plates were incubated for 60 min at 25 C. A diluent control consisting of block buffer (50 μΐ) also was placed in duplicate wells. Washing three times with PBST, 50 μΐ of biotinlabeled MAb 4.23 diluted 1:160 in block buffer were added to each well and incubated for 60 min at 25 C. Washing three times with PBST, 75 μΐ of streptavidin-horseradish peroxidase (Kirkegaard and Perry Laboratories) diluted 1:200 in block buffer was added to each well and incubated for 30 min at 37 C. Washing three times with PBST, 100 μΐ of 2,2’-azino-bis (3- ethylbenzthiazoline-6-sulfonic acid) substrate (Kirkegaard and Perry Laboratories) were added to each well; color development was stopped after 20 min with 1% (w/v) SDS in water. Plates were read on an ELISA reader (BT 2000 MicroKinetics Reader, Fisher Scientific) at 405 nm. Optical densities of duplicate wells, including positive and negative control sera wells, and diluent wells were averaged. Percentage of inhibition of optical densities of test serum wells relative to negative control serum was calculated after subtracting the diluent control, which was subtracted from all test and control well averages, to yield a corrected value. Percentage of inhibition was calculated as 100 - (100 x [test serum - diluent/negative control - diluent]). Sera were considered to be positive if inhibition >45% was observed and negative if inhibition